Inflammasome activation in myelodysplastic syndromes

ABSTRACT

Disclosed are methods for diagnosing a myelodysplastic syndrome (MDS) in a subject. In some embodiments, the method involves assaying a sample from the subject to detect inflammasome activation, wherein an increase in inflammasome activation in the sample compared to a control is an indication of MDS in the subject. The disclosed methods can further involve treating the subject for MDS if an increase in inflammasome activation is detected.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application is a continuation of copending application Ser. No.15/570,019, filed Nov. 26, 2019, which was the National Stage ofInternational Application No. PCT/US2016/030244, filed Apr. 29, 2016,which claims benefit of U.S. Provisional Application No. 62/155,994,filed May 1, 2015, and U.S. Provisional Application Ser. No. 62/307,960,filed Mar. 14, 2016, which are hereby incorporated herein by referencein their entirety.

STATEMENT REGARDING FEDERALLY SPONSORED RESEARCH OR DEVELOPMENT

This invention was made with Government Support under Grant No. CA115308and Grant No. CA187020 awarded by the National Institutes of Health. TheGovernment has certain rights in the invention.

BACKGROUND

Myelodysplastic syndromes (MDS) are hematopoietic stem cell malignanciescharacterized by dysplastic and ineffective hematopoiesis. MDS bonemarrow precursors have a larger cell size, deregulated proliferation andmaturation, and accelerated attrition by programmed cell death (List A,et al. N Engl J Med. 2005 352(6):549-57; Span L. F, et al. Leuk Res.2007 31(12):1659-67; Garcia-Manero G. Am J Hematol. 2014 89(1):97-108).Despite these shared phenotypes, MDS harbor a spectrum of clonalchromosome abnormalities and somatic gene mutations, the latter mostcommonly involving genes encoding RNA splicing and epigenetic regulatoryproteins (Bejar R, et al. J Clin Oncol. 2011 29(5):504-15; Yoshida K, etal. Nature. 2011 478(7367):64-9). How such diverse genetic eventsinitiate a common MDS phenotype is unexplained.

SUMMARY

Despite remarkable genetic heterogeneity, myelodysplastic syndromes(MDS) share features of cytological dysplasia and ineffectivehematopoiesis. Activation of the NLRP3 inflammasome that drives clonalexpansion and pyroptotic cell death is shown herein to be a hallmark ofMDS hematopoietic stem/progenitor cells (HSPC). Independent of genotype,MDS HSPC overexpress inflammasome proteins and manifest activated NLRP3complexes with consequent caspase-1 activation, IL-1β and IL-18generation, and pyroptotic cell death. Mechanistically, pyroptosis wastriggered by the alarmin S100A9 found in excess in MDS HSPC and bonemarrow plasma, which like founder gene mutations, induced reactiveoxygen species (ROS) to initiate cation influx, cell swelling andβ-catenin activation. Accordingly, knockdown of caspase-1,neutralization of S100A9 in BM plasma, and pharmacologic inhibition ofthe NLRP3 inflammasome or NADPH oxidase suppressed pyroptosis, ROSgeneration and nuclear β-catenin in MDS HSPC while restoring effectivehematopoiesis. Thus, DAMP signals and oncogenic mutations in MDS HSPClicense a common redox-sensitive inflammasome platform to inducepyroptosis and self-renewal, suggesting new avenues for therapeuticintervention that restore effective hematopoiesis in MDS patients.

Therefore, disclosed are methods for diagnosing a myelodysplasticsyndrome (MDS) in a subject. For example, the subject can be a patientsuspected of having a hematological disorder. The median age atdiagnosis of a MDS is between 60 and 75 years; a few patients areyounger than 50. Therefore, in some embodiments, the subject is at least50, 55, or 60 years of age. Signs and symptoms of MDS include anemia,neutropenia, and thrombocytopenia.

Also disclosed is a method for diagnosing Cryopyrin-Associated PeriodicSyndromes (CAPS) in a subject. There are 3 subtypes of CAPS: FamilialCold Autoinflammatory Syndrome (FCAS), Muckle-Wells Syndrome (MWS), andNeonatal-Onset Multisystem Inflammatory Disease (NOMID). CAPS aregenerally caused by autosomal-dominant mutations of the NLRP3 gene andresultant alterations in the protein cryopyrin, which NLRP3 encodes.Autoinflammatory diseases have symptoms that may resemble those ofautoimmune disorders, but they also have distinct clinicalpathophysiological features. The underlying diseases are characterizedby an inflammatory reaction that is seemingly unprovoked. Recurrentepisodes occur with clinical manifestations that vary by disease. Unlikeautoimmune diseases, autoinflammatory diseases are not associated withhigh-titer autoantibodies or antigen-specific T cells.

Also disclosed is a method for diagnosing an autoimmune disorder in asubject. Autoimmune diseases arise from an abnormal immune response ofthe body against substances and tissues normally present in the body(autoimmunity). The most common autoimmune disorders include rheumatoidarthritis, Lupus, Celiac disease, Sjögren's syndrome, Polymyalgiarheumatic, Multiple sclerosis, Ankylosing spondylitis, Type 1 diabetes,Alopecia areata, Vasculitis, and Temporal arteritis.

In some embodiments, the method involves assaying a sample from thesubject to detect inflammasome activation, wherein an increase ininflammasome activation in the sample compared to a control is anindication of MDS, CAPS, or autoimmune disorder in the subject.

An inflammasome is a multiprotein oligomer of caspase 1, PYCARD/ASC, anda member of the NOD-like receptor (NLR) family. NLRP1, NLRP3 and NLRC4are subsets of the NLR family and thus have two common features: thefirst is a nucleotide-binding domain (NBD) which is bound to byribonucleotide-phosphates (rNTP) and is important forself-oligomerization. The second is a C-terminus leucine-rich repeat(LRR), which serves as a ligand-recognition domain for other receptors(e.g. TLR) or microbial ligands. In some embodiments, the inflammasomeis an NLRP3 inflammasome.

Inflammasome activation can detected directly or indirectly. Forexample, inflammasome activation can be detected by assaying for geneexpression of CASP-1 (caspase-1 gene), NLRP3, or a combination thereof.Inflammasome activation can be detected by assaying for proteinexpression of active caspase-1, NLPR3, or a combination thereof.Inflammasome activation can also be detected by assaying directly forNLRP3 inflammasome complexes. For example, inflammasome activation canbe detected by detecting co-localization of active caspase-1 and NLRP3protein. Inflammasome activation can detected by detecting apoptosisassociated speck-like protein containing a CARD (ASC) specks.

The inflammasome promotes the maturation of the inflammatory cytokinesInterleukin 1β(IL-1β) and Interleukin 18 (IL-18). Therefore, in someembodiments, the method involves detecting IL-1β levels, IL-18, levels,or a combination thereof.

The sample of the disclosed methods preferably comprises hematopoieticstem/progenitor cells (HSPC). Therefore, in some embodiments, the sampleis a bone marrow sample.

The details of one or more embodiments of the invention are set forth inthe accompanying drawings and the description below. Other features,objects, and advantages of the invention will be apparent from thedescription and drawings, and from the claims.

DESCRIPTION OF DRAWINGS

FIGS. 1A to 1H show fulminant pyroptosis is manifest in HSPC and theirprogeny in MDS. FIG. 1A is a series of bar graphs showing markedlyincreased expression of pyroptosis-associated genes in MNC isolated frompatient MDS BM specimens (n=10 total; n=5 lower and n=5 higher-riskdisease), compared to normal controls (n=5). FIG. 1B is a representativeconfocal fluorescence micrograph (2520× magnification, 7.5 μm scale) ofa-caspase-1 and NLRP3 expression in MDS versus normal BM-MNC. DAPI,a-caspase-1, NLRP3; merged image shows inflammasome formation. FIG. 1Cshows quantitative analysis of a-caspase-1/NLRP3 confocal images ofBM-MNC isolated from lower-risk (n=7) and higher-risk (n=3) MDSpatients, and normal donors (n=6). MDS patient specimens havesignificantly increased expression of a-caspase-1 and NLRP3 proteins,accompanied by co-localization, confirming inflammasome assembly. FIG.1D contains representative scatter plots of pyroptotic cells(a-caspase-1⁺/annexin-V⁺) by flow cytometry in four phenotypicallydistinct hematopoietic lineages and cell types: stem cells (CD34⁺CD38⁻),progenitor cells (CD34⁺CD38⁺), immature myeloids (CD33⁺), and erythroids(CD71⁺). FIG. 1E shows quantitation of the mean percentage of pyroptoticcells by hematopoietic lineage in MDS (n=8) versus normal donors (n=5).FIG. 1F shows comparison of the mean percentage of pyroptotic versusapoptotic cells (a-caspase-3/7⁺/annexin-V⁺) by hematopoietic lineage inlower-risk MDS specimens (n=5). FIGS. 1G and 1H show mean percentage ofpyroptoic cells when lower-risk MDS BM-MNC (n=3) were transfected bylentivirus with shRNAs targeting CASP1 (FIG. 1G) or CASP3 (FIG. 1H).Suppression of caspase-1 resulted in a significant reduction in thepercentage of pyroptotic cells, whereas suppression of caspase-3 had nosignificant effect, confirming caspase-1 dependence. Error bars: SE,*p<0.05, ** p<0.01, and ***p<0.001.

FIGS. 2A to 2G show S100A9 initiates pyroptosis in MDS. FIG. 2A showsELISA assessment of BM plasma concentration of S100A9 in normal donors(n=12) versus lower-risk (n=33) and higher-risk (n=27) MDS. FIG. 2Bshows BM plasma concentration of HMGB1 assessed by ELISA in normaldonors (n=11) and MDS (n=55). FIG. 2C shows qPCR analysis of S100A9 mRNAin normal (n=2) versus lower-risk MDS (n=8). FIG. 2D shows HMGB1transcript levels in normal (n=6) versus MDS BM-MNC (n=10). FIG. 2Eshows changes in pyroptosis-related gene expression assessed by qPCR inuntreated normal BM-MNC (n=3), normal BM-MNC treated with 1 μg/mLrhS100A9 for 24 hours (n=2), and in MDS patient specimens (n=5). FIG. 2Fshows representative micrograph (2520× magnification, 7.5 μm scale)depicting inflammasome formation in normal, untreated BM-MNC or normalBM-MNC treated with 5 μg/mL rhS100A9 for 24 hours. DAPI a-caspase-1, andNLRP3; merged images show formation of inflammasome complexes. FIG. 2Gshows quantitative analysis of a-caspase-1/NLRP3 confocal images fromnormal donors (n=6), normal BM-MNC treated with 5 μg/mL rhS100A9 (n=2),and MDS patients (n=10). Error bars: SE, *p<0.05, ** p<0.01, and***p<0.001.

FIGS. 3A to 3J show inflammasome-initiated pore formation increases sizeof MDS precursors. FIG. 3A shows mean cell area quantified followinganalysis of a-caspase-1/NLRP3 confocal images of BM from normal donor(n=6), lower-risk (n=7), and higher-risk MDS patient specimens (n=3).Mean cell area is augmented in MDS irrespective of risk score. FIG. 3Bshows NLRP3 MFI and cell area are correlated in lower-risk MDS patients(r=0.49, n=7). FIG. 3C shows U937 cells incubated with 12.5 μg/mLethidium bromide (EB), and treated with 5 μg/mL rhS100A9 or media alone.Uptake of EB is depicted in green (680× magnification, 25 μm scale).FIG. 3D shows BM-MNC from normal donors (n=3) and MDS patients (n=3)incubated with autologous BM plasma for 24 hours. 12.5 μg/mL EB was thenadded to the cells, and dye incorporation measured by flow cytometry at5 minute intervals. FIG. 3E shows Left to right, photomicrograph imagesfrom normal donors illustrating normal red blood cell (RBC, 7.0 μm)followed by normal erythroid lineage maturation of nucleated BMprecursors with corresponding cell diameter. FIG. 3F shows correspondingimages from MDS BM aspirates, demonstrating an oval macrocyte (RBC, 9.1μm) followed by dysplastic and megaloblastic erythroid lineagematuration. FIG. 3G shows normal myelocyte. FIG. 3H shows enlargeddysplastic myelocyte with mild hypogranulation in MDS. FIGS. 3I and 3Hshow comparison of mean cell diameter in normal (n=4) versus MDS (n=4)BM during erythroid (FIG. 3I) and myeloid (FIG. 3J) lineage maturation.Left to right, maturation is depicted as most to least mature cellpopulations. Error bars: SE, *p<0.05, ** p<0.01, and ***p<0.001.

FIGS. 4A to 4E show inhibition of pyroptosis abrogates MDS HSPC celldeath and augments colony forming capacity. FIG. 4A shows reduction inthe fraction of pyroptotic BM cells from a MDS patient followingtreatment with 0.5 μg CD33-IgG₁ or 0.1 μM IRAK4 inhibitor. Values arenormalized to autologous BM plasma-incubated MDS BM-MNC. FIG. 4B showsquantitation of the mean percentage of pyroptotic cells in eachrespective lineage in MDS BM-MNC incubated with autologous BM plasma andeither 0.5 μg CD33-IgG₁ or 0.1 μM IRAK4 inhibitor for 24 hours (n=4).FIG. 4C shows BM-MNC isolated from lower-risk MDS patients (n=5) weretreated for 24 hours with CD33-IgG₁, and pyroptosis-related geneexpression was assessed by qPCR. FIGS. 4D and 4E shows colony formingcapacity was assessed in BM-MNC from MDS patient specimens (n=3) thatwere treated with increasing concentrations of CD33-IgG₁ (FIG. 4D), orwith the inflammasome inhibitor MCC950 (FIG. 4E). Error bars: SE.

FIGS. 5A to 5H show pyroptosis is the principal mechanism of HSPC deathin S100A9 transgenic mice. FIG. 5A shows quantitative analysis ofa-caspase-1 MFI, NLRP3 MFI, co-localization, and cell area from confocalimages of BM cells isolated from WT (n=2), 2 month (n=4), 6 month (n=5),and 11 month (n=4) old S100A9Tg mice. FIG. 5B shows representativemicrograph (2520× magnification, 7.5 μm scale) of a-caspase-1 and NLRP3protein expression in WT BM cells treated for 24 hours with 5 μg/mLS100A9, and of BM cells from S100A9Tg mice. DAPI, a-caspase-1, andNLRP3; merged image illustrates inflammasome formation. FIG. 5C showsquantitative analysis of confocal images of BM cells isolated from WT(n=2) mice, from WT BM cells treated for 24 hours with 5 μg/mL S100A9,or from BM cells from S100A9Tg mice (n=13). FIGS. 5D and 5E showrepresentative scatter plots of pyroptotic (FIG. 5D) and apoptotic (FIG.5E) KLS (c-Kit⁺Lin-Sca-1⁺) cells isolated from WT and transgenic mice.FIG. 5F shows mean percentage of pyroptotic versus apoptotic KLS cellsin WT (n=6) and S100A9Tg mice (n=6). FIG. 5G shows mean percentage oftotal a-caspase-1⁺ and a-caspase-3/7⁺ KLS cells isolated from WT (n=6)and S100A9Tg mice (n=6). FIG. 5H shows at six months of age, S100A9Tgtransgenic mice treated every other day with 50 mg/kg of theinflammasome inhibitor ICTA by oral gavage for a total of eight weeks.Shown are changes in hemoglobin, white blood cells (WBC), RBC andplatelet counts in WT (n=4) and S100A9Tg (n=5) versus S100A9Tg micetreated with ICTA (n=5). Error bars: SE, *p<0.05, ** p<0.01, and***p<0.001.

FIGS. 6A to 6H show S100A9 and MDS gene mutations induce ROS throughNADPH oxidase to activate β-catenin. FIG. 6A shows the percentage of ROSpositive cells. FIG. 6B shows ROS MFI assessed by flow cytometry inBM-MNC isolated from MDS patients (n=5) and normal donors (n=2). FIG. 6Cshows representative micrograph (2520× magnification, 7.5 μm scale) ofβ-catenin expression in normal BM-MNC (n=3), normal BM-MNC treated with5 μg/mL rhS100A9 (n=3) and MDS patients (n=6). DAPI and β-catenin areindividually depicted, and the merged image illustrates nuclearlocalization of β-catenin. FIG. 6D shows quantitation and scoring ofβ-catenin confocal images based on the presence of no, low, medium, orhigh nuclear β-catenin in normal BM-MNC, rhS100A9 treated normal BMcells, and MDS patients. FIG. 6E shows mean percentage of ROS positivecells and FIG. 6F shows ROS MFI assessed by flow cytometry in U2AF1 S34Fmutant expressing TF-1 cells and corresponding WT cells. Data arerepresentative of three independent experiments. FIG. 6G showsrepresentative micrograph (1890× magnification, 10 μm scale)illustrating β-catenin expression in U2AF1 WT cells, cells expressingS34F, or S34F-expressing mutant cells treated with NAC or the NADPHoxidase inhibitor DPI for 24 hours prior to staining. FIG. 6H showsquantitation and scoring of β-catenin confocal images based on thepresence of no, low, medium, or high nuclear β-catenin. Error bars: SE,*p<0.05, ** p<0.01, and ***p<0.001.

FIGS. 7A to 7C show caspase-1 activation exceeds caspase-3 activation inMDS. FIGS. 7A to 7C are bar graph showing mean percentage of totala-caspase-1⁺ (FIG. 7A), annexin-V⁺ (FIG. 7B), and a-caspase-3/7⁺ (FIG.7C) cells assessed by hematopoietic lineage. Data are representative offive normal donors and eight lower-risk MDS patients. Error bars: SE,*p<0.05 and ** p<0.01.

FIGS. 8A and 8B show gene expression following lentivirus-mediatedtransfection of MDS BM-MNC. Gene expression of CASP1 (FIG. 8A) and CASP3(FIG. 8B) was assessed by qPCR following shRNA-directed silencingperformed by lentivirus transfection of lower-risk BM-MNC (n=3).

FIGS. 9A to 9F show S100A9 provokes pyroptosis and inflammasomeactivation ex vivo. FIG. 9A shows U937 monocytic cells were treated withincreasing concentrations of rhS100A9 for 24 hours, resulting in aconcentration-dependent increase in the fraction of pyroptotic cells.FIG. 9B shows active caspase-1 MFI and percent positive cells increasein a concentration-dependent manner. FIG. 9C shows U937 cells treatedwith 5 μg/mL rhS100A9 overtime show a time-dependent increase ina-caspase-1 MFI and percent positive cells. FIG. 9D shows representativehistogram depicting a-caspase-1. LPS was used as a positive control forcaspase-1 activation. FIG. 9E shows representative micrograph (1890×magnification, 10 μm scale) depicting inflammasome formation in U937cells that were untreated or treated with 5 μg/mL rhS100A9 for 24 hours.DAPI, a-caspase-1, NLRP3; merged image shows formation of inflammasomecomplexes. FIG. 9F shows quantitative analysis of a-caspase-1/NLRP3confocal images of untreated and treated U937 cells. Cells were pooledfor analysis. Error bars: SE, ** p<0.01 and ***p<0.001. Data arerepresentative of three independent experiments.

FIG. 10 shows plasma levels of S100A9 are elevated in MDS. BM plasmaconcentration of S100A9 was assessed by ELISA and analyzed according toIPSS risk score. Only MDS patients with lower-risk disease (IPSS=0,IPSS=1) demonstrate a statistically significant increase in BM plasmaS100A9 concentration (p=2.3×10⁻⁵ and 1.0×10⁻³, respectively). Errorbars: SE, ***p<0.001.

FIGS. 11A to 11C show intracellular levels of the alarmin S100A9 areincreased across myeloid lineages. Intracellular levels of S100A9 weremeasured by flow cytometry in BM-MNC isolated from MDS patients (n=6)and normal controls (n=4). FIG. 11A is representative histogram. FIG.11B shows mean percentage of S100A9⁺ cells and FIG. 11C shows S100A9 MF.Error bars: SE, *p<0.05.

FIGS. 12A to 12E show increased size of MDS hematopoietic lineage cells.Images depict Wright-Giemsa staining (1000× magnification). FIG. 12Ashows normal BM with mild erythroid hyperplasia. The erythroidprecursors show a full spectrum of maturation with mean cellulardiameter recorded at different maturation stages [orthrochromicnormoblast, 7.5 μm (normal reference: 6-12 μm); early to latepolychromic normoblasts, 8.5 μm (normal reference: 8-14 μm); early andlate basophilic normoblasts, 12.4 μm (normal reference: 12-17 μm and10-15 μm, respectively); and promonoblasts, 15.8 μm (normal reference:14-24 μm)]. FIG. 12B shows dysplastic erythroid precursors in the BMfrom an MDS patient. The erythroid precursors show obscured stagespecific maturation or maturation asynchrony. Hematopoietic precursorsare enlarged in size compared to the corresponding stage of maturationin normal donors [dysplastic/megaloblastoid orthrochromic normoblasts(15.8 μm), dysplastic early to late polychromic binucleated normoblast(18.2 μm), dysplastic late basophilic normoblasts (17.6 μm), anddysplastic promonoblasts (25.5 μm)]. FIG. 12C shows normal BM withcomplete spectrum of myeloid maturation. The myeloid progenitorsrepresent different stages of maturation with appropriate size[segmented neutrophil (10-18 μm), band form (10-20 μm), metamyelocyte(10-18 μm), myelocyte (10-20 μm), promyelocyte (12-24 μm), andmyeloblasts (9-20 μm)]. FIG. 12D shows enlarged eosinophilic myelocytesmeasuring 23.1 μm at the maximal dimension in a background of markeddyserythropoiesis. FIG. 12E shows enlarged myelocyte with overtmaturation asynchrony in a background of dyserythropoiesis in an MDS BM.The myelocyte measures 23.6 μm, which is larger than a normal myelocyte.

FIGS. 13A and 13B show ICTA inhibits inflammasome activation. FIG. 13Ais a representative micrograph (1890× magnification) depictinginflammasome formation in U937 cells following 24 hour treatment withvehicle or 5 μg/mL rhS100A9 alone or with ICTA (20 μg/mL). DAPI,a-caspase-1, NLRP3; merged image shows formation of inflammasomecomplexes. FIG. 13B shows quantitative analysis of confocal images.Error bars: SE, *p<0.05, **p<0.01, ***p<0.001.

FIGS. 14A to 14D show recombinant human S100A9 is sufficient to induceROS and β-catenin activation in monocytic cells. U937 cells were treatedwith 5 μg/mL rhS100A9 for 24 hours. FIGS. 14A and 14B show thepercentage of ROS positive cells was assessed by flow cytometry (FIG.14A) and DNA damage assessed by comet assay (FIG. 14B). FIG. 14C showsrepresentative micrograph of β-catenin (1890× magnification, 10 μmscale) in untreated and treated cells by confocal microscopy. DAPI(blue), β-catenin; merged image shows nuclear localization of β-catenin,reflecting its active form. FIG. 14D shows β-catenin confocal imageswere quantified and scored based on the presence of no, low, medium, orhigh expression of nuclear β-catenin. Cells were pooled for analysis.There is a statistically significant increase in high nuclear β-cateninexpression following treatment with rhS100A9 (p=2.4×10⁻³). Error bars:SE, **p<0.01 and ***p<0.001. Data are representative of threeindependent experiments.

FIGS. 15A to 15J show U2AF1 mutations manifest in MDS provokepyroptosis. The ability of U2AF1 mutations to induce pyroptosis wasassessed in S34F mutant cell lines. FIG. 15A shows representativedensity plot of inflammasome formation based on the detection offluorescence pulse differences in ASC. FIG. 15B shows quantitation ofASC in WT, S34F, and S34F cells treated with DPI for 24 hours. FIG. 15Cshows representative scatter plots of pyroptotic cells by flowcytometry. FIG. 15D shows quantitation of the percentage of pyroptoticcells in mutant and WT cells. FIGS. 15E to 14H show the relativepercentage of total a-caspase-1⁺ (FIG. 15E) and annexin-V⁺ cells (FIG.15F), as well as the MFI of a-caspase-1 (FIG. 15G) and annexin-V (FIG.15H) assessed by flow cytometry. FIG. 15I shows Mean cell area wasquantitated from confocal images of WT and S34F mutant cells. FIG. 15Jshows that to investigate pore formation, 12.5 μg/mL EB was added to theWT and mutant line, and incorporation of the dye was measured by flowcytometry at 5 min intervals. Error bars: SE, *p<0.05, ** p<0.01, and***p<0.001. Data are representative of three independent experiments.

FIGS. 16A to 16G shows SF3B1 K700E induces pyroptosis. The ability ofthe SF3B1 K700E conditional knock-in mutation to induce pyroptosis wasassessed in BM cells harvested from WT (n=3) and mutant (n=3) mice. FIG.16A shows quantitation of the percentage of pyroptotic versus apoptoticcells. FIG. 16B shows mean percentage of total a-caspase-1⁺,a-caspase-3/7⁺, and annexin-V⁺ cells. FIG. 16C shows MFI values fora-caspase-1, a-caspase-3/7, and annexin-V in the mutant and WT cells.FIG. 16D shows representative micrograph (2520× magnification, 7.5 μmscale) depicting inflammasome formation in the WT and K700E mutantcells. DAPI, a-caspase-1, NLRP3; merged image shows inflammasomeformation. FIG. 16E shows quantitative analysis of a-caspase-1/NLRP3confocal images. FIG. 16F shows representative density plot ofinflammasome formation based on the detection of fluorescence pulsedifferences in ASC. FIG. 16G shows quantitation of ASC in WT (n=6),K700E (n=6), and K700E cells treated with NAC (n=6) or DPI (n=3) for 24hours.

FIGS. 17A to 17D show SF3B1 K700E supports self-renewal throughβ-catenin activation. The ability of the SF3B1 K700E conditionalknock-in mutation to support self-renewal through activation ofβ-catenin was assessed in BM cells harvested from WT (n=6) and mutant(n=6) mice. FIG. 17A shows mean percentage of ROS-positive cells, andFIG. 17B shows ROS MFI assessed by flow cytometry. FIG. 17C showsrepresentative micrograph (2520× magnification, 7.5 μm scale) ofβ-catenin expression. DAPI, β-catenin, and the merged images shownuclear localization of β-catenin. FIG. 17D shows quantitation andscoring of β-catenin confocal images based on the presence of no, low,medium, or high nuclear β-catenin in WT (n=6), K700E (n=6), and K700Ecells treated with NAC (n=3) or DPI (n=3) for 24 hours. Error bars: SE,*p<0.05 and ** p<0.01.

FIGS. 18A to 18K show SRSF2 mutants induce pyroptosis and supportself-renewal through β-catenin. HEK293T cells were transientlytransfected with WT and P95H mutant SRSF2. Data are representative ofthree independent experiments, and of the GFP⁺ transfected population.FIG. 18A shows representative density plot of inflammasome formationbased on the detection of fluorescence pulse differences in ASC. FIG.18B shows quantitation of ASC positive cells. FIGS. 18C to 18G show foldchange of the mean percentage of pyroptotic cells (FIG. 18C), totala-caspase-1⁺ cells (FIG. 18D), total annexin-V⁺ cells (FIG. 18E), andMFI values for a-caspase-1 (FIG. 18F) and annexin-V (FIG. 18G),normalized to WT transfected cells. FIG. 18H shows mean percentage ofROS positive cells and FIG. 18I shows ROS MFI. FIG. 18J showsrepresentative micrograph (1890× magnification, 10 μm scale) ofβ-catenin expression. DAPI, β-catenin, and the merged images shownuclear localization of β-catenin. FIG. 18K shows quantitation andscoring of β-catenin confocal images based on the presence of no, low,medium, or high nuclear β-catenin. Error bars: SE, *p<0.05.

FIGS. 19A to 19H show Asxl1 and Tet2 deletions are sufficient to inducepyroptosis. The ability of deletions in Asxl1 and Tet2 to inducepyroptosis was assessed in BM cells isolated from Asxl1 KO, Tet2 KO, andDKO cells, compared to control cells. FIG. 19A shows representativemicrograph (2520× magnification, 7.5 μm scale) depicting inflammasomeformation in control and KO cells that were untreated, or treated withNAC or DPI. DAPI, a-caspase-1, NLRP3; merged images show inflammasomeformation. FIGS. 19B to 19D show quantitative analysis ofa-caspase-1/NLRP3 confocal images. Cells were pooled for analysis. FIG.19E shows representative density plot of inflammasome formation based onthe detection of fluorescence pulse differences in ASC. FIG. 19F showsquantitation of percentage of ASC positive cells. FIG. 19G showsquantitation of mean cell area. FIG. 19H shows to assess pore formation,ethidium bromide was added to the cells and dye incorporation wasmeasured by flow cytometry at 5 minute intervals. Error bars: SE,*p<0.05, ** p<0.01, and ***p<0.001.

FIG. 20A to 20D show Asxl1 and Tet2 deletions are sufficient to driveself-renewal through β-catenin activation. The ability of deletions inAsxl1 and Tet2 to activate β-catenin was assessed in BM cells isolatedfrom Asxl1 KO, Tet2 KO, and DKO cells, compared to control cells. FIG.20A shows mean percentage of ROS positive cells, and FIG. 20B shows ROSMFI assessed by flow cytometry. FIG. 20C shows representative micrograph(2520× magnification, 7.5 μm scale) of β-catenin expression. DAPI,β-catenin, and the merged images show nuclear localization of β-catenin.FIG. 20D shows quantitation and scoring of β-catenin confocal imagesbased on the presence of no, low, medium, or high nuclear β-catenin.Cells were pooled for analysis, and measurements of significance weremade on untreated deleted cells compared to NAC or DPI treated deletedcells. Error bars: SE, *p<0.05 and ***p<0.001.

FIG. 21 shows a S100A9-pyroptosis circuit provokes phenotypes manifestin MDS. a) S100A8/A9 binds both CD33 and TLR4, resulting in inflammasomeassembly. Ligation of S100A8/A9 to TLR4 results in NFκB-mediatedtranscription and subsequent production of proinflammatory cytokinessuch as pro-IL-1β and pro-IL-18, along with inflammasome components. b)through interaction with Rac2 and p67phox, S100A8/A9 promotes activationof NOX, which results in a dual function. First, NOX proteins generateROS, which serve to activate NLRs and inflammasome assembly. Second,NOX-derived ROS results in oxidation of NRX, leading to its dissociationfrom Dvl. Once dissociated, Dvl suppress the β-catenin destructioncomplex (GSKβ/CK1/APC/Axin), resulting in stabilization of β-catenin.This allows β-catenin to enter the nucleus and induce transcription ofTCF/LEF controlled genes, including cyclin-D1 and c-Myc, which areessential to self-renewal. c) transient receptor potential melastatin 2(TRPM2), a calcium-permeable cation channel in hematopoietic cells, isactivated by NOX-derived ROS via oxidation of a single channelmethionine residue, Met²¹⁴. Upon activation, TRPM2 causes an influx ofcalcium leading to mitochondrial depolarization and further release ofROS, which activate the inflammasome complex. d) formation of theinflammasome complex occurs as a consequence of ROS activation and DAMPsignaling. Once activated, inflammasomes mediate conversion ofpro-caspase-1 to its mature and catalytically active form. Activecaspase-1 cleaves pro-IL-1β and pro-IL-18 to their mature forms. e)pyroptosis ensues with loss of membrane integrity resulting in releaseof pro-inflammatory cytokines and other intracellular contents into theextracellular milieu. f) MDS-related gene mutations activate NF-κB andNLRP3 via NOX-generated ROS (Sallmyr A, et al. Cancer Lett. 2008270(1):1-9; Rassool F, et al. Cancer Res. 2007 67(18):8762-71).

FIGS. 22A and 22B are plots showing percentage (FIG. 22A) and MFI (FIG.22B) of ASC specks in hematologic malignancies, including chronicmyelomonocytic leukemia (CMML, n=20), chronic lymphocytic leukemia (CLL,n=50) and acute myeloid leukemia (AML, n=10).

DETAILED DESCRIPTION

Apoptosis, a non-inflammatory form of programmed cell death, has beenimplicated in the ineffective hematopoiesis in MDS based upon membraneexternalization of phosphatidylserine, mitochondrial depolarization andDNA fragmentation (Raza A, et al. Blood. 1995 86(1):268-76; Parker J. E,et al. Blood. 2000 96(12):3932-8; Tehranchi R, et al. Blood. 2003101(3):1080-6). Nevertheless, the inflammatory cytokine and cellularmilieu instead support innate immune activation (Takizawa H, et al.Blood 2012 119:2991-3002). Inflammatory cytokines such as interleukin-1β(IL-1β), tumor necrosis factor-α, transforming growth factor-β, IL-6 andothers are generated in excess in MDS, accompanied by bone marrowexpansion of hematopoietic-inhibitory, myeloid derived suppressor cells(MDSC) activated by the danger associated molecular pattern (DAMP)S100A9, a Toll-like receptor (TLR)-4/CD33 ligand (Mundle S. D, et al.Blood. 1996 88(7):2640-7; Chen X, et al. J Clin Invest. 2013123(11):4595-611; Vogl T, et al. Nat Med. 2007 13(9):1042-9; Ehrchen J.M, et al. J Leukoc Biol. 2009 86(3):557-66). MDS hematopoietic stem andprogenitor cells (HSPC) express elevated levels of TLRs and theirsignaling intermediates, while TLR signaling has been implicated in theproliferation of MDS HSPC and in the pathogenesis of peripheral bloodcytopenias (Maratheftis C. I., et al. Clin Cancer Res. 200713(4):1154-60; Rhyasen G. W, et al. Cancer Cell. 2013 24(1):90-104;Hofmann W. K, et al. Blood. 2002 100(10):3553-60).

Recent studies have shown that activation of TLRs by select DAMPs cantrigger pyroptosis, a novel caspase-1-dependent pro-inflammatory celldeath (Masters S. L, et al. Immunity. 2012 37(6):1009-23; Brennan M. A.,et al. Mol Microbiol. 2000 38(1):31-40; Cookson B. T. & Brennan M. A.Trends Microbiol. 2001 9(3):113-4) that involves the activation of iongradients, cell swelling and the release of IL-1β and IL-18,intracellular DAMPs and other pro-inflammatory cytokines (Ehrchen J. M,et al. J Leukoc Biol. 2009 86(3):557-66; Maratheftis C. I., et al. ClinCancer Res. 2007 13(4):1154-60; Rhyasen G. W, et al. Cancer Cell. 201324(1):90-104; Masters S. L, et al. Immunity. 2012 37(6):1009-23;Bergsbaken T, et al. Nat Rev Microbiol. 2009 7(2):99-109). Pyroptosis ismediated by the formation of inflammasome complexes, cytosolicheptameric oligomers composed of nucleotide-binding domain andleucine-rich repeat pattern recognition receptors (NLRs). The bestcharacterized NLR, NLRP3, undergoes a conformational change in responseto DAMP interaction to recruit the adapter protein, apoptosis-associatedspeck-like protein containing a caspase-recruitment domain (ASC), andpro-caspase-1, which in turn catalyzes proteolytic cleavage of pro-IL-1β& pro-IL-18 to their active forms (Brennan M. A., et al. Mol Microbiol.2000 38(1):31-40). Inflammasome activation involves NFκB-inducedtranscriptional priming of inflammasome components, followed by cationchannel activation with cell volume expansion and inflammasome componentassembly (Brennan M. A., et al. Mol Microbiol. 2000 38(1):31-40; CooksonB. T. & Brennan M. A. Trends Microbiol. 2001 9(3):113-4; Bergsbaken T,et al. Nat Rev Microbiol. 2009 7(2):99-109; Fantuzzi G. & Dinarello, C.A. J Clin Immunol. 1999 19(1):1-11). Inflammasome assembly is induced byS100A9 homodimers and S100A8 heterodimers, which function as alarminsregulating NADPH oxidase to generate reactive oxygen species (ROS), andwhich extracellularly direct paracrine inflammatory signals (Kessel C,et al. Clin Immunol. 2013 147(3):229-41; Lim S. Y., et al. J LeukocBiol. 2009 86(3):577-87; Simard J. C, et al. PLoS One. 20138(8):e72138).

S100A9 and ROS, generated in response to NLRP3 activation or somaticgene mutations, are shown herein to serve as DAMP signalingintermediates responsible for inflammasome-mediated pyroptosis andβ-catenin activation in MDS. Further, targeting this circuit restoreseffective hematopoiesis in MDS. Collectively, these features account forthe key biological features of the MDS phenotype and suggest novelstrategies for therapeutic intervention.

All inflammasomes require the adapter protein apoptosis associatedspeck-like protein containing a CARD (ASC) for the activation ofcaspase-1. After inflammasome activation, ASC assembles into a largeprotein complex, which is termed “speck”. ASC specks can be observed asthey reach a size of around 1 μm and in most cells only one speck formsupon inflammasome activation. Therefore, in some embodiments, ASC speckformation can be used as an upstream readout for inflammasomeactivation.

In some aspects, the disclosed method is an immunoassay. Immunoassays,in their most simple and direct sense, are binding assays involvingbinding between antibodies and antigen. For example, antibodies thatspecifically bind human a-caspase-1, NLPR3, IL-1β, IL-18, and S100A9 arecommercially available and can be produced using routine skill.

Many types and formats of immunoassays are known and all are suitablefor detecting the disclosed biomarkers. Examples of immunoassays areenzyme linked immunosorbent assays (ELISAs), radioimmunoassays (RIA),radioimmune precipitation assays (RIPA), immunobead capture assays,Western blotting, dot blotting, gel-shift assays, Flow cytometry,protein arrays, multiplexed bead arrays, magnetic capture, in vivoimaging, fluorescence resonance energy transfer (FRET), and fluorescencerecovery/localization after photobleaching (FRAP/FLAP).

In general, immunoassays involve contacting a sample suspected ofcontaining a molecule of interest (such as the disclosed biomarkers)with an antibody to the molecule of interest or contacting an antibodyto a molecule of interest (such as antibodies to the disclosedbiomarkers) with a molecule that can be bound by the antibody, as thecase may be, under conditions effective to allow the formation ofimmunocomplexes. Contacting a sample with the antibody to the moleculeof interest or with the molecule that can be bound by an antibody to themolecule of interest under conditions effective and for a period of timesufficient to allow the formation of immune complexes (primary immunecomplexes) is generally a matter of simply bringing into contact themolecule or antibody and the sample and incubating the mixture for aperiod of time long enough for the antibodies to form immune complexeswith, i.e., to bind to, any molecules (e.g., antigens) present to whichthe antibodies can bind. In many forms of immunoassay, thesample-antibody composition, such as a tissue section, ELISA plate, dotblot or Western blot, can then be washed to remove any non-specificallybound antibody species, allowing only those antibodies specificallybound within the primary immune complexes to be detected.

Immunoassays can include methods for detecting or quantifying the amountof a molecule of interest (such as the disclosed biomarkers or theirantibodies) in a sample, which methods generally involve the detectionor quantitation of any immune complexes formed during the bindingprocess. In general, the detection of immunocomplex formation is wellknown in the art and can be achieved through the application of numerousapproaches. These methods are generally based upon the detection of alabel or marker, such as any radioactive, fluorescent, biological orenzymatic tags or any other known label. See, for example, U.S. Pat.Nos. 3,817,837; 3,850,752; 3,939,350; 3,996,345; 4,277,437; 4,275,149and 4,366,241, each of which is incorporated herein by reference in itsentirety and specifically for teachings regarding immunodetectionmethods and labels.

As used herein, a label can include a fluorescent dye, a member of abinding pair, such as biotin/streptavidin, a metal (e.g., gold), or anepitope tag that can specifically interact with a molecule that can bedetected, such as by producing a colored substrate or fluorescence.Substances suitable for detectably labeling proteins include fluorescentdyes (also known herein as fluorochromes and fluorophores) and enzymesthat react with colorometric substrates (e.g., horseradish peroxidase).The use of fluorescent dyes is generally preferred in the practice ofthe invention as they can be detected at very low amounts. Furthermore,in the case where multiple antigens are reacted with a single array,each antigen can be labeled with a distinct fluorescent compound forsimultaneous detection. Labeled spots on the array are detected using afluorimeter, the presence of a signal indicating an antigen bound to aspecific antibody.

A modifier unit such as a radionuclide can be incorporated into orattached directly to any of the compounds described herein byhalogenation. In another aspect, the radionuclide can be attached to alinking group or bound by a chelating group, which is then attached tothe compound directly or by means of a linker. Radiolabeling techniquessuch as these are routinely used in the radiopharmaceutical industry.

Labeling can be either direct or indirect. In direct labeling, thedetecting antibody (the antibody for the molecule of interest) ordetecting molecule (the molecule that can be bound by an antibody to themolecule of interest) include a label. Detection of the label indicatesthe presence of the detecting antibody or detecting molecule, which inturn indicates the presence of the molecule of interest or of anantibody to the molecule of interest, respectively. In indirectlabeling, an additional molecule or moiety is brought into contact with,or generated at the site of, the immunocomplex. For example, asignal-generating molecule or moiety such as an enzyme can be attachedto or associated with the detecting antibody or detecting molecule. Thesignal-generating molecule can then generate a detectable signal at thesite of the immunocomplex. For example, an enzyme, when supplied withsuitable substrate, can produce a visible or detectable product at thesite of the immunocomplex. ELISAs use this type of indirect labeling.

As another example of indirect labeling, an additional molecule (whichcan be referred to as a binding agent) that can bind to either themolecule of interest or to the antibody (primary antibody) to themolecule of interest, such as a second antibody to the primary antibody,can be contacted with the immunocomplex. The additional molecule canhave a label or signal-generating molecule or moiety. The additionalmolecule can be an antibody, which can thus be termed a secondaryantibody. Binding of a secondary antibody to the primary antibody canform a so-called sandwich with the first (or primary) antibody and themolecule of interest. The immune complexes can be contacted with thelabeled, secondary antibody under conditions effective and for a periodof time sufficient to allow the formation of secondary immune complexes.The secondary immune complexes can then be generally washed to removeany non-specifically bound labeled secondary antibodies, and theremaining label in the secondary immune complexes can then be detected.The additional molecule can also be or include one of a pair ofmolecules or moieties that can bind to each other, such as thebiotin/avadin pair. In this mode, the detecting antibody or detectingmolecule should include the other member of the pair.

Other modes of indirect labeling include the detection of primary immunecomplexes by a two step approach. For example, a molecule (which can bereferred to as a first binding agent), such as an antibody, that hasbinding affinity for the molecule of interest or corresponding antibodycan be used to form secondary immune complexes, as described above.After washing, the secondary immune complexes can be contacted withanother molecule (which can be referred to as a second binding agent)that has binding affinity for the first binding agent, again underconditions effective and for a period of time sufficient to allow theformation of immune complexes (thus forming tertiary immune complexes).The second binding agent can be linked to a detectable label orsignal-generating molecule or moiety, allowing detection of the tertiaryimmune complexes thus formed. This system can provide for signalamplification.

Immunoassays that involve the detection of as substance, such as aprotein or an antibody to a specific protein, include label-free assays,protein separation methods (i.e., electrophoresis), solid supportcapture assays, or in vivo detection. Label-free assays are generallydiagnostic means of determining the presence or absence of a specificprotein, or an antibody to a specific protein, in a sample. Proteinseparation methods are additionally useful for evaluating physicalproperties of the protein, such as size or net charge. Capture assaysare generally more useful for quantitatively evaluating theconcentration of a specific protein, or antibody to a specific protein,in a sample. Finally, in vivo detection is useful for evaluating thespatial expression patterns of the substance, i.e., where the substancecan be found in a subject, tissue or cell.

In some aspects, the method comprises detecting gene expression forgenes such as CASP-1, NLRP3, or a combination thereof, e.g., using aprimer or probe that selectively binds CASP-1 or NLRP3 mRNA.

A number of widely used procedures exist for detecting and determiningthe abundance of a particular mRNA in a total or poly(A) RNA sample. Forexample, specific mRNAs can be detected using Northern blot analysis,nuclease protection assays (NPA), in situ hybridization, or reversetranscription-polymerase chain reaction (RT-PCR).

In theory, each of these techniques can be used to detect specific RNAsand to precisely determine their expression level. In general, Northernanalysis is the only method that provides information about transcriptsize, whereas NPAs are the easiest way to simultaneously examinemultiple messages. In situ hybridization is used to localize expressionof a particular gene within a tissue or cell type, and RT-PCR is themost sensitive method for detecting and quantitating gene expression.

Northern analysis presents several advantages over the other techniques.The most compelling of these is that it is the easiest method fordetermining transcript size, and for identifying alternatively splicedtranscripts and multigene family members. It can also be used todirectly compare the relative abundance of a given message between allthe samples on a blot. The Northern blotting procedure isstraightforward and provides opportunities to evaluate progress atvarious points (e.g., intactness of the RNA sample and how efficientlyit has transferred to the membrane). RNA samples are first separated bysize via electrophoresis in an agarose gel under denaturing conditions.The RNA is then transferred to a membrane, crosslinked and hybridizedwith a labeled probe. Nonisotopic or high specific activity radiolabeledprobes can be used including random-primed, nick-translated, orPCR-generated DNA probes, in vitro transcribed RNA probes, andoligonucleotides. Additionally, sequences with only partial homology(e.g., cDNA from a different species or genomic DNA fragments that mightcontain an exon) may be used as probes.

The Nuclease Protection Assay (NPA) (including both ribonucleaseprotection assays and S1 nuclease assays) is an extremely sensitivemethod for the detection and quantitation of specific mRNAs. The basisof the NPA is solution hybridization of an antisense probe (radiolabeledor nonisotopic) to an RNA sample. After hybridization, single-stranded,unhybridized probe and RNA are degraded by nucleases. The remainingprotected fragments are separated on an acrylamide gel. Solutionhybridization is typically more efficient than membrane-basedhybridization, and it can accommodate up to 100 μg of sample RNA,compared with the 20-30 μg maximum of blot hybridizations. NPAs are alsoless sensitive to RNA sample degradation than Northern analysis sincecleavage is only detected in the region of overlap with the probe(probes are usually about 100-400 bases in length).

NPAs are the method of choice for the simultaneous detection of severalRNA species. During solution hybridization and subsequent analysis,individual probe/target interactions are completely independent of oneanother. Thus, several RNA targets and appropriate controls can beassayed simultaneously (up to twelve have been used in the samereaction), provided that the individual probes are of different lengths.NPAs are also commonly used to precisely map mRNA termini andintron/exon junctions.

In situ hybridization (ISH) is a powerful and versatile tool for thelocalization of specific mRNAs in cells or tissues. Unlike Northernanalysis and nuclease protection assays, ISH does not require theisolation or electrophoretic separation of RNA. Hybridization of theprobe takes place within the cell or tissue. Since cellular structure ismaintained throughout the procedure, ISH provides information about thelocation of mRNA within the tissue sample.

The procedure begins by fixing samples in neutral-buffered formalin, andembedding the tissue in paraffin. The samples are then sliced into thinsections and mounted onto microscope slides. (Alternatively, tissue canbe sectioned frozen and post-fixed in paraformaldehyde.) After a seriesof washes to dewax and rehydrate the sections, a Proteinase K digestionis performed to increase probe accessibility, and a labeled probe isthen hybridized to the sample sections. Radiolabeled probes arevisualized with liquid film dried onto the slides, while nonisotopicallylabeled probes are conveniently detected with colorimetric orfluorescent reagents.

RT-PCR has revolutionized the study of gene expression. It is nowtheoretically possible to detect the RNA transcript of any gene,regardless of the scarcity of the starting material or relativeabundance of the specific mRNA. In RT-PCR, an RNA template is copiedinto a complementary DNA (cDNA) using a retroviral reversetranscriptase. The cDNA is then amplified exponentially by PCR. As withNPAs, RT-PCR is somewhat tolerant of degraded RNA. As long as the RNA isintact within the region spanned by the primers, the target will beamplified.

Relative quantitative RT-PCR involves amplifying an internal controlsimultaneously with the gene of interest. The internal control is usedto normalize the samples. Once normalized, direct comparisons ofrelative abundance of a specific mRNA can be made across the samples. Itis crucial to choose an internal control with a constant level ofexpression across all experimental samples (i.e., not affected byexperimental treatment). Commonly used internal controls (e.g., GAPDH,β-actin, cyclophilin) often vary in expression and, therefore, may notbe appropriate internal controls. Additionally, most common internalcontrols are expressed at much higher levels than the mRNA beingstudied. For relative RT-PCR results to be meaningful, all products ofthe PCR reaction must be analyzed in the linear range of amplification.This becomes difficult for transcripts of widely different levels ofabundance.

Competitive RT-PCR is used for absolute quantitation. This techniqueinvolves designing, synthesizing, and accurately quantitating acompetitor RNA that can be distinguished from the endogenous target by asmall difference in size or sequence. Known amounts of the competitorRNA are added to experimental samples and RT-PCR is performed. Signalsfrom the endogenous target are compared with signals from the competitorto determine the amount of target present in the sample.

As disclosed herein, the disclosed indications of inflammasomeactivation and s100A9 levels can also be used to predict whether thesubject has low-risk or high-risk MDS, CAPS, or autoimmune disorder.Therefore, this can be used to select the appropriate therapy, dosage,or combination thereof.

MDSs are hematological (blood-related) medical conditions withineffective production (or dysplasia) of the myeloid class of bloodcells. In some cases, the MDS patient has a chromosome 5q deletion(del(5q)). However, in other cases, the patient has non-del5q MDS.

In some embodiments, MDS can be treated with a therapeutically effectiveamount of inflammasome inhibitor. Inflammasome inhibitors are describedin PCT/US2016/019925, which is incorporated by reference for theteaching of these inhibitors and their use in treating an MDS.

In some embodiments, the method involves treating the subject with atherapeutically effective amount of lenalidomide.

In some cases, the myeloid disorder is amyelodysplastic/myeloproliferative neoplasms (MDS/MPN). In some cases,the myeloid disorder is a myelodysplastic syndrome withmyeloproliferative features. In some cases, the myeloid disorder is atherapy-related myeloid neoplasm.

Cryopyrin-associated periodic syndrome (CAPS) is a spectrum ofautoinflammatory syndromes including familial cold autoinflammatorysyndrome (FCAS, formerly termed familial cold-induced urticaria), theMuckle-Wells syndrome (MWS), and neonatal-onset multisystem inflammatorydisease (NOMID, also called chronic infantile neurologic cutaneous andarticular syndrome or CINCA). They share many clinical features. Thesesyndromes are associated with mutations in NLRP3, the gene encodingcryopyrin. This is a component of the interleukin 1 inflammasome, andmutations lead to unregulated production of interleukin 1β. Monoclonalantibodies against interleukin 1β (such as canakinumab), otherinterleukin 1 binding proteins (such as rilonacept), or interleukin 1receptor antagonist (for example anakinra) can be used to treat thesedisorders.

Autoimmune diseases arise from an abnormal immune response of the bodyagainst substances and tissues normally present in the body(autoimmunity). The most common autoimmune disorders include rheumatoidarthritis, Lupus, Celiac disease, Sjögren's syndrome, Polymyalgiarheumatic, Multiple sclerosis, Ankylosing spondylitis, Type 1 diabetes,Alopecia areata, Vasculitis, and Temporal arteritis. The treatment ofautoimmune diseases is typically with immunosuppression. Treatmentsinclude Cytokine blockade (or the blockade of cytokine signalingpathways), removal of effector T-cells and B-cells (e.g. anti-CD20therapy can be effective at removing instigating B-cells), andintravenous immunoglobulin.

The disclosed compositions, including pharmaceutical composition, may beadministered in a number of ways depending on whether local or systemictreatment is desired, and on the area to be treated. For example, thedisclosed compositions can be administered intravenously,intraperitoneally, intramuscularly, subcutaneously, intracavity, ortransdermally. The compositions may be administered orally, parenterally(e.g., intravenously), by intramuscular injection, by intraperitonealinjection, transdermally, extracorporeally, ophthalmically, vaginally,rectally, intranasally, topically or the like, including topicalintranasal administration or administration by inhalant.

Parenteral administration of the composition, if used, is generallycharacterized by injection. Injectables can be prepared in conventionalforms, either as liquid solutions or suspensions, solid forms suitablefor solution of suspension in liquid prior to injection, or asemulsions. A revised approach for parenteral administration involves useof a slow release or sustained release system such that a constantdosage is maintained.

The compositions disclosed herein may be administered prophylacticallyto patients or subjects who are at risk for MDS. Thus, the method canfurther comprise identifying a subject at risk for MDS prior toadministration of the herein disclosed compositions.

The exact amount of the compositions required will vary from subject tosubject, depending on the species, age, weight and general condition ofthe subject, the severity of the allergic disorder being treated, theparticular nucleic acid or vector used, its mode of administration andthe like. Thus, it is not possible to specify an exact amount for everycomposition. However, an appropriate amount can be determined by one ofordinary skill in the art using only routine experimentation given theteachings herein. For example, effective dosages and schedules foradministering the compositions may be determined empirically, and makingsuch determinations is within the skill in the art. The dosage rangesfor the administration of the compositions are those large enough toproduce the desired effect in which the symptoms disorder are affected.The dosage should not be so large as to cause adverse side effects, suchas unwanted cross-reactions, anaphylactic reactions, and the like.Generally, the dosage will vary with the age, condition, sex and extentof the disease in the patient, route of administration, or whether otherdrugs are included in the regimen, and can be determined by one of skillin the art. The dosage can be adjusted by the individual physician inthe event of any counterindications. Dosage can vary, and can beadministered in one or more dose administrations daily, for one orseveral days. Guidance can be found in the literature for appropriatedosages for given classes of pharmaceutical products. A typical dailydosage of the disclosed composition used alone might range from about 1μg/kg to up to 100 mg/kg of body weight or more per day, depending onthe factors mentioned above.

In some embodiments, the molecule containing lenalidomide isadministered in a dose equivalent to parenteral administration of about0.1 ng to about 100 g per kg of body weight, about 10 ng to about 50 gper kg of body weight, about 100 ng to about 1 g per kg of body weight,from about 1 μg to about 100 mg per kg of body weight, from about 1 μgto about 50 mg per kg of body weight, from about 1 mg to about 500 mgper kg of body weight; and from about 1 mg to about 50 mg per kg of bodyweight. Alternatively, the amount of molecule containing lenalidomideadministered to achieve a therapeutic effective dose is about 0.1 ng, 1ng, 10 ng, 100 ng, 1 μg, 10 μg, 100 μg, 1 mg, 2 mg, 3 mg, 4 mg, 5 mg, 6mg, 7 mg, 8 mg, 9 mg, 10 mg, 11 mg, 12 mg, 13 mg, 14 mg, 15 mg, 16 mg,17 mg, 18 mg, 19 mg, 20 mg, 30 mg, 40 mg, 50 mg, 60 mg, 70 mg, 80 mg, 90mg, 100 mg, 500 mg per kg of body weight or greater.

The term “subject” refers to any individual who is the target ofadministration or treatment. The subject can be a vertebrate, forexample, a mammal. Thus, the subject can be a human or veterinarypatient. The term “patient” refers to a subject under the treatment of aclinician, e.g., physician.

The term “therapeutically effective” refers to the amount of thecomposition used is of sufficient quantity to ameliorate one or morecauses or symptoms of a disease or disorder. Such amelioration onlyrequires a reduction or alteration, not necessarily elimination.

The term “sample from a subject” refers to a tissue (e.g., tissuebiopsy), organ, cell (including a cell maintained in culture), celllysate (or lysate fraction), biomolecule derived from a cell or cellularmaterial (e.g. a polypeptide or nucleic acid), or body fluid from asubject. Non-limiting examples of body fluids include blood, urine,plasma, serum, tears, lymph, bile, cerebrospinal fluid, interstitialfluid, aqueous or vitreous humor, colostrum, sputum, amniotic fluid,saliva, anal and vaginal secretions, perspiration, semen, transudate,exudate, and synovial fluid.

The term “treatment” refers to the medical management of a patient withthe intent to cure, ameliorate, stabilize, or prevent a disease,pathological condition, or disorder. This term includes activetreatment, that is, treatment directed specifically toward theimprovement of a disease, pathological condition, or disorder, and alsoincludes causal treatment, that is, treatment directed toward removal ofthe cause of the associated disease, pathological condition, ordisorder. In addition, this term includes palliative treatment, that is,treatment designed for the relief of symptoms rather than the curing ofthe disease, pathological condition, or disorder; preventativetreatment, that is, treatment directed to minimizing or partially orcompletely inhibiting the development of the associated disease,pathological condition, or disorder; and supportive treatment, that is,treatment employed to supplement another specific therapy directedtoward the improvement of the associated disease, pathologicalcondition, or disorder.

A number of embodiments of the invention have been described.Nevertheless, it will be understood that various modifications may bemade without departing from the spirit and scope of the invention.Accordingly, other embodiments are within the scope of the followingclaims.

EXAMPLES Example 1: Identification of the NLRP3 Inflammasome as a Driverof the MDS Phenotype

Methods

MDS patient specimens. MDS patients consented on The University of SouthFlorida Institutional Review Board approved protocols were recruitedfrom the Malignant Hematology Clinic at H. Lee Moffitt Cancer Center &Research Institute, and the Eastern Cooperative Oncology Group (ECOG)E2905 trial (NCT00843882). Pathologic subtype of MDS was reportedaccording to World Health Organization (WHO) criteria and prognosticrisk assigned according to the International Prognostic Scoring System(IPSS). Patients were segregated as lower (Low, Intermediate-1) andhigher risk (Intermediate-2, High) MDS. Bone marrow mononuclear cells(BM-MNC) were isolated from heparinized bone marrow aspirates usingFicoll-Hypaque Plus gradient centrifugation (GE Healthcare).

Mice. S100A9Tg mice were used for animal studies (Chen X, et al. J ClinInvest. 2013 123(11):4595-611). WT FVB/NJ mice were purchased fromJackson Laboratories (Bar Harbor, Me.). Heparinized BM cells wereisolated from tibias and femurs of male and female mice.

Reagents and cells. U937 cells were grown in RPM11640 supplemented with10% FBS. TF-1 U2AF1 mutant and mock WT cells were cultured in RPM11640supplemented with 10% FBS and 2 ng/mL recombinant human GM-CSF. Cellswere maintained at 37° C. under 5% CO₂. Normal, heparinized BM aspirateswere purchased from Lonza Walkersville or AllCells, LLC. Recombinanthuman S100A9 and the CD33/Siglec 3 chimeric fusion protein weregenerated as previously described (Chen X, et al. J Clin Invest. 2013123(11):4595-611). The IRAK4 inhibitor was acquired through materialtransfer agreement from Nimbus Discovery. NAC and DPI were purchasedfrom Sigma-Aldrich. Active caspase-1 and caspase-3/7 were detected usingFAM-FLICA® Caspase-1 and Caspase-3/7 activity kits, (ImmunoChemistryTechnologies). NLRP1 antibodies were purchased from Santa CruzBiotechnology, NLRP3 antibodies from Abcam, and β-catenin antibodiesfrom BD Biosciences.

Pyroptosis flow cytometry panel. For human samples, treated anduntreated BM-MNC were incubated overnight in IMDM, supplemented with 10%autologous BM plasma. Subsequently, cells were harvested, washed twicein 1× PBS, and stained with LIVE/DEAD Violet fluorescent reactive dyeaccording to the manufacturer's protocol (Life Technologies). Cells wereresuspended in 1× PBS with 2% BSA, and incubated at room temperature for15 minutes to block non-specific binding. After washing, cells werestained with 30× FAM-FLICA® Caspase-1 and Caspase-3/7 solution at aratio of 1:30 for 2 hours at 37° C. Cells were washed and stained forcell surface receptors using CD38:PE-CF594, CD33:BV711, CD34:APC (BDBiosciences), and CD71:PE-Cyanine7 (eBioscience). All antibodies werediluted 1:20, and cells were stained for 30 minutes at 4° C. Cells werewashed, resuspended in 1× binding buffer, and stained withAnnexin-V:Cy5.5 at a dilution of 1:20 for 15 minutes at room temperature(BD Biosciences). 1× binding buffer was added to a final volume of 400μL. Sample acquisitions were carried out using a BD LSR II flowcytometer and FACSDiva software (BD Biosciences). Calibration of theflow cytometer was carried out prior to each experiment using RainbowMid-Range Fluorescent Particles (BD Biosciences). To establishfluorescence compensation settings, ArC Amine Reactive CompensationBeads were used for LIVE/DEAD Violet staining (Life Technologies), andBD CompBead Plus Anti-Mouse Ig κ/Negative Control (BSA) CompensationPlus Particles were used for surface receptor conjugates (BDBiosciences). Data were analyzed using FlowJo 9.7.5 software (FlowJo,LLC, Ashland, Oreg.).

S100A9 flow cytometry experiments in U937 cells. Monocytic U937 cellswere treated with the indicated concentrations of rhS100A9 for 24 hours,or with 5 μg/mL rhS100A9 for the indicated time points. Subsequently,cells were stained with 30×FAM-FLICA® Caspase-1 solution at a ratio of1:30 for 2 hours at 37° C. Cells were washed, resuspended in 1× bindingbuffer, and stained with Annexin-V:PE at a dilution of 1:30 for 15minutes at room temperature. 1× binding buffer was added to a finalvolume of 350 μL. Sample acquisitions were carried out using a BDFACSCalibur flow cytometer (BD Biosciences). Data were analyzed usingFlowJo 9.7.5 software.

Intracellular S100A9 flow cytometry. BM-MNC were incubated overnight inIMDM, supplemented with 10% autologous BM plasma. The following day,cells were harvested and washed twice in 1× PBS. Cells were fixed withBD Cytofix Fixation Buffer at 37° C. for 10 minutes, and stored at −80°C. until staining. At the time of staining, cells were warmed to 37° C.in a water bath, spun down, and washed 1× with staining buffer. Pelletswere resuspended in 1 mL of pre-warmed BD Permeabilization Buffer III,and incubated on ice for 30 minutes. Cells were washed twice withstaining buffer. Following washing, cells were stained with S100A9:FITC(BioLegend), and cell surface receptors using CD38:PE-CF594, CD33:BV711,CD34:APC (BD Biosciences), and CD71:PE-Cyanine7 (eBioscience). Allantibodies were diluted 1:20, and cells were stained for 30 minutes at4° C. Cells were washed and resuspended in 400 μL staining buffer.Sample acquisitions were carried out using a BD LSR II flow cytometerand FACSDiva software (BD Biosciences).

Enzyme-linked immunosorbent assays (ELISA). Human S100A9 DuoSet ELISAkit was purchased from R&D Systems and HMGB1 ELISA kit was purchasedfrom MYBioSource. Both were performed according to manufacturer'sprotocol.

Immunofluorescence confocal microscopy. MDS and normal donor BM-MNC andmouse BM cells were stained with 30× FAM-FLICA® Caspase-1 solution at aratio of 1:30 for 2 hours at 37° C. Cells were washed and cytospins weregenerated using a 5 minutes centrifugation at 450 rpm. Slides were fixedat 37° C. for 10 minutes using BD Cytofix Fixation Buffer (BDBiosciences), and subsequently washed using PBS. Cells werepermeabilized with 0.1% Triton X-100/2% BSA in PBS for 15 minutes atroom temperature. After washing with PBS, cells were blocked using 2%BSA in PBS for 30 minutes at room temperature, and washed again. Cellswere incubated with the appropriate primary antibody overnight (1:400for NLRP3, 1:20 for β-catenin) at 4° C. The next day, cells were washedwith PBS and incubated with the appropriate secondary antibodies (1:500)for 1 hour at room temperature. After washing, cells were covered withProLong Gold Antifade Reagent with DAPI prior to the addition of acoverslip (Life Technologies). Co-localization of a-caspase-1 with NLRP3inflammasome complexes was assessed using a Leica TCS SP5 AOBS LaserScanning Confocal microscope (Leica Microsystems). Analysis of theinflammasome images was performed with Definiens Developer 2.0(Definiens AG). The software was used to segment cells based onbrightness and size thresholds, followed by a watershed segmentationalgorithm. Intensity values and Pearson's correlation coefficient wereextracted from the segmented cells. For β-catenin image analysis,confocal images were imported into Definiens Tissue Studio v3.0, 64 Dualin .tif format. Cells were separated from background using the RGBthresholds. Nuclei were identified by setting thresholds in the DAPIchannel. Typical cells averaged 60 microns. The red intensity(β-catenin) in the nucleus and cytoplasm was established by settingthresholds to low, medium and high in the red channel on a scale of0-255 in the red channel.

ASC staining to detect inflammasome formation by flow cytometry.Staining was carried out as described (Sester D. P, et al. J Immunol.2015 194(1):455-62). Briefly, cell pellets were resuspended in 1 mL ofprewarmed BD Permeabilization Buffer III, and incubated on ice for 30minutes. Cells were washed 2× with staining buffer. Following washing,cells were stained with rabbit-anti-ASC primary antibodies at a 1:1500dilution and incubated for 90 minutes. Cells were washed, stained withsecondary antibodies at a dilution of 1:1500, and incubated for 45minutes. Cells were washed, and sample acquisitions were carried outusing a BD LSR II flow cytometer and FACSDiva software.

Real-time quantitative PCR. RNA was isolated from BM-MNC using theRNeasy Mini Kit (Qiagen). cDNA was produced using the iScript cDNASynthesis Kit (Bio-Rad). Sequences for primers can be found in Table 1.GAPDH mRNA was used for transcript normalization. cDNA was amplifiedusing the iQ SYBR Green Supermix and the CFX96 Real-Time PCR DetectionSystem (Bio-Rad). PCR conditions were as follows: 10 minutes at 95° C.,followed by 40 cycles of amplification (15 seconds at 95° C. and 1minute at 60° C.). Relative gene expression was calculated using the−2^(ΔΔCt) method.

TABLE 1 Primer sets used for qPCR. Gene Forward CASP15′-TGAGCAGCCAGATGGTAGAGC-3′ SEQ ID NO: 1 CASP35′-GTGAGGCGGTTGTAGAAGAGTTTC-3′ SEQ ID NO: 2 IL-1β5′-CTCTTCGAGGCACAAGGCAC-3′ SEQ ID NO: 3 IL-18 5′-ACTGCCTGGACAGTCAGCAA-3′SEQ ID NO: 4 NLRP1 5′-TCTACGTTGGCCACTTGGGA-3′ SEQ ID NO: 5 NLRP35′-CAATGGGGAGGAGAAGGCGT-3′ SEQ ID NO: 6 S100A95′-CTCGGCTTTGACAGAGTGCAA-3′ SEQ ID NO: 7 HMGB15′-CCCTCCCAAAGGGGAGACAAA-3′ SEQ ID NO: 8 GAPDH 5′-GAAGGTGAAGGTCGGACT-3′SEQ ID NO: 9 Reverse CASP1 5′-TCACTTCCTGCCCACAGACAT-3′ SEQ ID NO: 10CASP3 5′-TGAGCAGGGCTCGCTAACTC-3′ SEQ ID NO: 11 IL-1β5′-CAAGTCATCCTCATTGCCACTGT-3′ SEQ ID NO: 12 IL-185′-GCAGCCATCTTTATTCCTGAGA-3′ SEQ ID NO: 13 NLRP15′-AGAGGTGAAGGTACGGCTATGC-3′ SEQ ID NO: 14 NLRP35′-TCTGAACCCCACTTCGGCTC-3′ SEQ ID NO: 15 S100A95′-CTGGTTCAGGGTGTCTGGGT-3′ SEQ ID NO: 16 HMGB15′-AGAGGAAGAAGGCCGAAGGAG-3′ SEQ ID NO: 17 GAPDH5′-GAAGATGGTGATGGGATTTC-3′ SEQ ID NO: 18

Lentiviral infection of primary mononuclear cells. Lentiviral constructswere purchased from Origene. Caspase-1 (TL305640), Caspase-3(TL305638b), and scrambled (TR30021) HuSH™ shRNA plasmids were amplifiedby transforming One Shot® Top10 competent cells (Life technologies)according to manufacturer's protocol. Single colonies were expanded andmini preps were performed using the Qiagen QIAprep® Mini Prep Kit.HEK293T cells were transfected by incubating 2600 ng of shRNA plasmid,30 μL Lipofectamine® 2000 (Invitrogen), 26 μL MISSION™ LentiviralPackaging Mix (Sigma Aldrich) in 500 μL of Opti-MEM®I (Lifetechnologies) for 15 minutes at room temperature. This mixture was thenadded to 70% confluent HEK293T cells with 4 mL Opti-MEM®I medium withoutserum in a 100 mm dish. Cells were incubated at 37° C. for 6 hours in ahumidified chamber, then 6 mL of DMEM (Mediatech, Manassas, Va.) with10% serum was added. In the morning, medium was changed and 10 mL freshDMEM was added. Virus was collected at 48 and 72 hours using 0.45 μmfilters. The concentration was determined to be at least 5×10⁵ IFU/mLusing Clontech Lenti-X Go Stix (Mountain View, Calif.) and stored at 4°C. Virus up to one week old was used for experiments. For primary cellinfection, 2.5 million cells were plated in a 100 mm dish with 1.25 mLfresh virus, 1.25 mL opti-MEM®I, and 8 μg/mL polybrene. Cells wereincubated overnight, then 5 mL of fresh IMDM (Mediatech) with 10% FBSwas added. RNA was isolated after 72 hours of infection using QiagenRNeasy Isolation Kit according to manufacturer's protocol and mRNAlevels were analyzed by qPCR using GAPDH mRNA levels as a control.Additionally at 72 hours, 10% v/v of autologous BM plasma was added tothe remaining cells. Twenty-four hours after plasma was added, cellswere stained with annexin-V, 7-AAD, and FAM-FLICA® Caspase-1 andanalyzed using a BD FACSCalibur flow cytometer.

Colony formation assay. Four replicates of 350,000 BM-MNC/mL wereresuspended in 10% autologous BM plasma and plated in MethoCultmethylcellulose medium (Stemcell Technologies) supplemented with 1% v/vpenicillin-streptomycin and 3 units/mL erythropoietin. CD33-IgG andMCC950 were added directly to the medium prior to plating. Colonies ofBFU-E, CFU-GM, and CFU-GEMM were scored using an inverted lightmicroscope fourteen days after plating.

SRSF2 transfection of HEK293T cells. HEK293T cells were transfected byincubating 4 μg of SRSF2 DNA with 10 μL Lipofectamine® 2000 in 100 μL ofOpti-MEM®I for 20 minutes at room temperature. This mixture was added to70% confluent HEK293T cells with 2 mL Opti-MEM®I medium without serum ina 6 well plate. Cells were incubated at 37° C. for 4 hours in ahumidified chamber, then medium was replaced with 2 mL of DMEM with 10%serum added. Treatment with NAC or DPI and subsequent analyses werecarried out 24 and 48 hours following transfection, respectively.

Pore formation assay. MDS and normal donor BM-MNC were incubated in 10%autologous BM plasma at 37° C. overnight. Cells were subsequently washedand resuspended in 1 mL of PBS. 12.5 μg/mL ethidium bromide (FisherScientific, Pittsburgh, Pa.) was added, and sample acquisitions wereacquired at the indicated time points using a BD FACSCalibur flowcytometer.

ROS detection. ROS were detected using CM-H₂DCFDA and CellROX® Deep RedReagent according to manufacturer's protocol (Life Technologies).

Comet assays. Monocytic U937 cells were treated with 5 μg/mL rhS100A9for 24 hr. DNA damage was measured by single cell gel electrophoresisusing alkaline CometAssay® according to manufacturer's protocol(Trevigen Inc.).

ICTA mouse treatment studies. ICTA was synthesized by the Drug DiscoveryCore Facility at H. Lee Moffitt Cancer Center & Research Institute. Sixmonth old transgenic mice (n=5) were dosed every other day with 50 mg/kgICTA by oral gavage, for a total of eight weeks.

Statistics. Data are expressed as means±standard error. Statisticalanalyses were carried out in Microsoft Excel using student's t-test,correlations using chi square for non-continuous variables and logisticregression for continuous variables were performed using IPSS softwarev22 (SPSS Inc., Chicago, Ill.), and *p values<0.05, ** p<0.01, and***p<0.01 were considered to be statistically significant.

Results

MDS HSPC Manifest Inflammasome Activation and Pyroptosis

To assess whether pyroptosis was primed in MDS, expression of genesencoding inflammasome proteins was evaluated in bone marrow (BM)mononuclear cells (BM-MNC) isolated from MDS patients (n=10) compared toage-matched normal controls (n=5). MDS specimens displayed markedup-regulation of inflammasomal transcripts (FIG. 1a ). For example,caspase-1 (CASP1) gene expression was increased 209-fold in MDS, whereascaspase-3 (CASP3), the canonical apoptotic caspase, was 40% lower in MDScompared to normal controls. Gene expression of NLRP3 was increased48.1-fold in MDS. Further, the expression of the inflammatory cytokinesIL-1β and IL-18 was increased 3.7-fold and 29.6-fold in lower-risk MDS(n=5) compared to normal controls (n=5), whereas higher-risk MDSspecimens demonstrated only 1.1-fold and 9.2-fold up-regulation (n=5),consistent with the known up-regulation of survival signals in higherrisk MDS. Confocal fluorescence microscopy confirmed selectiveactivation of NLRP3 inflammasome complexes in MDS specimens versusage-matched control BM-MNC, where there was co-localization andincreased active (a)-caspase-1 (MFI increased 3.7-fold in lower-risk[p=7.1×10⁻³] and 4.1-fold in higher-risk disease [p=6.0×10⁻³]) and NLRP3(MFI was increased 69.1-fold in lower-risk [p=0.013] and 68.2-fold inhigher-risk [p=5.1×10⁻³]) disease (FIG. 1b, 1c ). MDS specimens alsodisplayed significantly greater inflammasome assembly compared tocontrols, irrespective of IPSS risk group (FIG. 1c ). Specifically,NLRP3 inflammasome assembly was increased 2.9-fold in lower-risk(p=3.9×10⁻⁵) and 3.1-fold in higher-risk (p=7.1×10⁻⁵) patients.

To assess pyroptosis in MDS, the percentage of pyroptotic cells, definedas percentage of a-caspase-1⁺/annexin-V⁺ cells, was determined inphenotypically distinct hematopoietic lineages by flow cytometry. Normal(n=5) and lower-risk MDS BM-MNC (n=8) were incubated with autologous BMplasma for 24 hours prior to flow cytometry analysis. MDS HSPCdemonstrated markedly increased pyroptosis (FIG. 1d ), where thefraction of pyroptotic cells increased 150-fold in CD34⁺CD38⁻ stem cells(p=0.051), 22.8-fold in progenitor cells (CD34⁺CD38⁺), 13.0-fold inimmature myeloid cells (CD33⁺), and 6.8-fold in erythroid cells (CD71⁺,p=3.1×10⁻³), compared to normal controls (FIG. 1e ). Additionally, thepercentage of a-caspase-1⁺ cells was increased 15.6-fold inhematopoietic stem cells (p=0.032), 14.1-fold in progenitors, 12.1-foldin immature myeloid cells (p=0.012), and 10.1-fold in CD71⁺ cells(p=1.5×10⁻³) (FIG. 7a ). Overall, only the stem cell population had asignificant increase in total annexin-V⁺ cells (p=3.8×10⁻³) (FIG. 7b ).A-caspase-1 MFI directly correlated with NLRP3 MFI, inflammasomeassembly, and the percentage of pyroptotic stem cells. Notably, thelatter was directly associated with the percentage of a-caspase-1⁺ CD33myeloid progenitors. The extent of apoptosis (i.e.,a-caspase-3/7⁺/annexin-V⁺) was also evaluated in lower-risk MDSspecimens (n=5). Pyroptotic cells were 1.2-fold, 1.6-fold, 1.9-fold, and3.6-fold up-regulated in stem cells, progenitor cells, immaturemyeloids, and erythroid cells, compared to the apoptotic cell fraction(FIG. 1f ). No significant differences in a-caspase-3/7⁺ cells weredetected in any of the four hematopoietic lineages investigated (FIG. 7c). Lastly, to confirm that caspase-1 is essential for hematopoietic celldeath in MDS, shRNA-directed silencing of caspase-1 and caspase-3 wasperformed by lentivirus transfection of lower-risk BM-MNC (n=3) (FIG.8). Knockdown of caspase-1 significantly decreased the fraction ofpyroptotic cells, greater than 35% versus scrambled transfected controls(p=0.038) (FIG. 1g ). In contrast, knockdown of caspase-3 had nodiscernible effect (FIG. 1h ), confirming selectivecaspase-1-dependence.

The DAMP Protein S100A9 is a Primary Initiator of Pyroptosis

BM plasma concentrations of the alarmin S100A9 are profoundly increasedin MDS and stimulate the expansion of MDSC through ligation of itscognate receptors, TLR4 and CD33 (Chen X, et al. J Clin Invest. 2013123(11):4595-611). As NLRPs are sensors of DAMP signals, experimentswere conducted to determine if S100A9 triggers pyroptosis in MDS.Indeed, treatment of the monocytic cell line U937 with recombinant humanS100A9 (rhS100A9) resulted in a concentration-dependent increase in thefraction of pyroptotic cells (FIG. 9a ), with a corresponding increasein a-caspase-1 MFI and percentage of a-caspase-1⁺ cells (FIG. 9b ).Likewise, treatment with 5 μg/mL rhS100A9 provoked a time-dependentincrease in activation of caspase-1 (FIG. 9c, 9d ). Furthermore,treatment with rhS100A9 markedly increased levels of NLRP3 inflammasomecomplexes and this was accompanied by caspase-1 activation (FIG. 9e, 9f).

The BM plasma concentration of S100A9 was significantly higher inlower-risk patient specimens (n=33) compared to normal controls (n=12;p=1.5×10⁻⁴), with no difference in higher-risk patient specimens (n=27,FIG. 2a ). Analysis of S100A9 BM plasma concentration by InternationalPrognostic Scoring System (IPSS) risk category showed a 2.3- and2.2-fold increase in low risk (n=10, p=2.3×10⁻¹) and intermediate-I(n=23, p=1.0×10⁻³), compared to normal controls (n=12), with nosignificant differences among controls and intermediate-II (n=17) orhigh risk (n=10) disease (FIG. 10). Notably, BM S100A9 concentrationswere significantly higher in lower-risk versus higher-risk MDS (p=0.013)(FIG. 2a ). In addition, the BM plasma concentration of HMGB1, a nuclearDAMP and TLR4 ligand, was significantly increased in MDS (n=55) versusnormal controls (n=11) (p=2.6×10⁻³) (FIG. 2b ) (Velegraki M, et al.Haematologica. 2013 98(8):1206-15; Chirico V, et al. Eur J Pediatr. 2014173(9):1123-36). Moreover, S100A9 and HMGB1 gene expression wereup-regulated 104.5-fold and 1.5-fold in MDS, respectively, compared tonormal controls (FIG. 2c, 2d ), and flow cytometric analysis confirmed acorresponding increase in cellular expression of the S100A9 alarmin inMDS stem cells and progeny (FIG. 11).

To determine if S100A9 directly triggers pyroptosis in HSPC, normalBM-MNC (n=2) were treated with 1 μg/mL rhS100A9 and changes in geneexpression was assessed by qPCR. The expression of pyroptosis-associatedgenes was significantly up-regulated following addition of rhS100A9, forsome to levels that surpass those detected in MDS (n=5) (FIG. 2e ).Accordingly, a-caspase-1 and NLRP3 levels were induced followingtreatment of normal BM-MNC with 5 μg/mL rhS100A9 (FIG. 2f ), by 2.5-foldand 47.1-fold (p=0.064), respectively, as were formation of NLRP3inflammasomes, by 2.9-fold (p=3.1×10⁻⁴) (FIG. 2g ). Finally, althoughrhS100A9 treatment induced inflammasome assembly and caspase-1activation in normal controls, MDS patient specimens (n=10) displayedgreater activation and co-localization of these effectors.

Inflammasome-Initiated Pore Formation Increases Size of MDS Precursors

Cell swelling is a hallmark of pyroptosis. This occurs followingcaspase-1-mediated activation of plasma membrane cation channels, whichcompromises membrane integrity and disrupts osmolality (Fink S. L. &Cookson B. T. Cell Microbiol. 2006 8(11):1812-25). Confocal imageanalyses of MDS BM-MNC cells demonstrated significantly larger mean cellarea compared to normal controls (FIG. 3a ). This phenotype wasaccentuated in lower-risk MDS patients compared to normal controls(p=6.0×10⁻⁵), with no significant difference detected in higher-riskpatients. Further, there was a positive correlation between mean NLRP3MFI and mean cell area in lower-risk (but not higher-risk) MDS patients(r=0.49) (p=7.8×10⁻³) (FIG. 3b ). To assess pore formation, influx ofthe membrane-impermeable, cationic dye ethidium bromide was assessed byimmunofluorescence. As predicted, monocytic U937 cells treated withrhS100A9 demonstrated rapid and substantial uptake of ethidium bromide(FIG. 3c ). Further, flow cytometric analyses of ethidium bromide uptakedemonstrated that MDS specimens incubated with autologous bone marrowplasma had rapid and sustained elevated dye influx versus that of BM-MNCfrom normal donors, which was demonstrable as early as 20 minutes(p=0.041), and remained significant through 1 hour of dye exposure(p=0.014) (FIG. 3d ). Finally, analysis of normal and MDS bone marrowaspirate morphology confirmed the larger cell size by maturation stageand lineage in MDS (FIG. 3e-j , 12).

Inhibition of Pyroptosis Improves Hematopoiesis in MDS

To assess the role of S100A9 in the pyroptosis phenotype evidenced inMDS, experiments were conducted to assess the effects of a S100A9high-affinity chimeric (CD33-IgG₁) decoy receptor or of an IRAK4inhibitor on phenotypes manifest in BM-MNC (n=4) from MDS patientstreated with autologous BM plasma. Notably, treatment with CD33-IgG₁ orthe IRAK4 inhibitor led to marked reduction in the fraction ofpyroptotic cells across all lineages studied (FIG. 4a ). Overall,short-term incubation with the chimera or the IRAK4 inhibitor reducedthe fraction of pyroptotic cells, with corresponding maximumlineage-specific changes in stem cells (44% vs. 75%, respectively),progenitor cells (23% vs. 36%, respectively), CD33⁺ (68% vs. 55%,respectively) and CD71⁺ cells (64% vs. 55%, respectively) (FIG. 4b ).Short-term treatment with the chimeric receptor also significantlyreduced the MDSC fraction, suggesting that S100A9 neutralization impairsthe survival of MDSC. Consistent with this, the CD33 chimera reducedexpression of CASP1, IL-1β, IL-18, and NLRP3 versus autologous BM plasmaalone (n=5) (FIG. 4c ). CASP3 expression was also markedly reduced,which is consistent with caspase-3 being activated downstream ofcaspase-1 after late mitochondrial depolarization (Ali A, et al. JHematother Stem Cell Res. 1999 8(4):343-56). High concentrations of thechimera led to cross-linking of the IgG₁-Fc domains and aggregation thatmasked dose-dependent effects of S100A9 neutralization.

To test if S100A9 neutralization could improve hematopoiesis in MDS,colony forming capacity was assessed after plating of MDS BM-MNC inautologous BM plasma and increasing concentrations of CD33-IgG₁ (FIG. 4d) or of MCC950 (FIG. 4e ), a small molecule inhibitor of NLRP3 (Coll R.C, et al. Nat Med. 2015 21(3):248-55). Neutralization of S100A9 orinhibition of the NLRP3 inflammasome markedly improved colony-formingcapacity (up to 6.6-fold greater than controls). Thus, pyroptoticpathway inhibition abrogates MDS hematopoietic cell death and promoteseffective hematopoiesis.

S100A9 is Sufficient to Provoke HSPC Pyroptosis In Vivo

To assess whether forced expression of S100A9 was sufficient to inducepyroptosis in vivo, an S100A9 transgenic (S100A9Tg) mouse model wasinvestigated that phenocopies human MDS (Chen X, et al. J Clin Invest.2013 123(11):4595-611). Confocal fluorescence microscopy analyses of BMcells from the tibia and femurs of S100A9Tg versus wild type (WT) miceat 2 (n=4), 6 (n=4), and 11 (n=5) months of age established thata-caspase-1 levels selectively increased in an age-dependent manner inthe BM of S100A9 transgenics, with a 2.1-fold up-regulation at 2 months,2.4-fold at 6 months (p=3.3×10⁻³), and 2.5-fold at 11 months (p=0.010)versus WT mice. Similarly NLRP3 levels were increased in S100A9Tg mice,with a 21.1-fold up-regulation at 2 months (p=0.059), 25.6-fold at 6months (p=2.2×10⁻⁴), and 12.1-fold at 11 months (p=0.018) (FIG. 5a ).Accordingly, formation of NLRP3 inflammasome complexes was significantlyincreased in an age-dependent fashion, with 2.6-fold greaterco-localization in the 2 month old S100A9Tg transgenic mice (p=0.017),3.3-fold in the 6 month (p=1.0×10⁻⁶), and 3.2-fold in 11 month old mice(p=1.2×10⁻³) (FIG. 5a ). Though transgenic mice illustrate a markedincrease in mean cell area at each time point, there was no significantdifference in cell size between WT and transgenic mice.

To test if S100A9 triggers pyroptosis in mouse hematopoietic cells, BMcells isolated from WT mice were treated with 5 μg/mL rhS100A9 andinflammasome formation was assessed by confocal microscopy (FIG. 5b, 5c). As predicted, MFI of a-caspase-1 and NLRP3 were both significantlyincreased after rhS100A9 treatment (n=2) versus controls (n=2)(p=7.5×10⁻³ and 0.017, respectively). Notably, MFI values from rhS100A9treated BM cells of WT mice were comparable to those manifest in the BMcells of S100A9 transgenic mice (n=13) (FIG. 5c ), and rhS100A9treatment of WT BM cells was associated with a marked induction ofinflammasome complexes (p=0.023). To assess the extent of pyroptosisversus apoptosis in the corresponding mice, BM cells were isolated from7 month old WT (n=6) and 9 month old S100A9Tg mice (n=6). Activecaspase-1 and a-caspases-3/7 were assessed by flow cytometry in the KLS(c-Kit⁺Lin⁻Sca-1⁺) hematopoietic stem and progenitor cell population(FIG. 5d, 5e ). The mean percentage of pyroptotic KLS cells wassignificantly increased in the S100A9Tg animals versus WT mice(p=0.038), whereas there were no significant differences in the meanpercentage of apoptotic cells (FIG. 5f ). Additionally, the totalpercentage of a-caspase-1⁺ KLS cells was increased 2.7-fold in theS100A9Tg mice compared to WT mice (p=2.75×10⁻⁴), with no change in thetotal a-caspase-3/7⁺ KLS population (FIG. 5g ). Finally, to test if invivo inflammasome inhibition improves hematopoiesis in S100A9Tg miceanalogous to human MDS, aged S100A9Tg mice were treated with ICTA, anIcariin derivative that inhibits NLRP3 inflammasome activation (FIG.13), every other day for eight weeks. ICTA treated transgenic miceshowed marked improvement in peripheral blood counts, includingincreased hemoglobin, leukocyte count, red blood cells and plateletcounts (FIG. 5h ), indicating restored effective hematopoiesis. Thus,pyroptosis is the principal mechanism driving HSPC cell death and MDS inS100A9Tg mice.

S100A9 and MDS Gene Mutations Trigger Pyroptosis and β-CateninActivation Via ROS

ROS act as DAMP intermediates that activate the Wnt/β-catenin axis(Rharass T, et al. J Biol Chem. 2014 289(40):27937-51; Kajla S, et al.FASEB J. 2012 26(5):2049-59). Thus, ROS generated by either S100A9 orsomatic gene mutations that are manifest in MDS may contribute toself-renewal and clonal expansion via activation of pi-catenin.Monocytic U937 cells treated with 5 μg/mL rhS100A9 for 24 hoursdisplayed a 2.9-fold increase in the mean percentage of ROS-positivecells and a 4.1-fold increase in DNA damage (p=1.5×10⁻⁹), compared tountreated cells (FIG. 14a, 14b ). Accordingly, mean percentage of ROSpositive cells was increased in MDS BM-MNC (n=5) 16.5-fold compared tonormal controls (n=2) (p=0.011) (FIG. 6a ), with a correspondingsignificant increase in ROS MFI (p=0.028) (FIG. 6b ). Further, confocalfluorescence microscopy analyses demonstrated marked increases in thelevels of nuclear (activated) β-catenin in rhS100A9 treated U937 cellscompared to untreated cells (p=2.4×10⁻³) (FIG. 14c, 14d ), and elevatedlevels of nuclear β-catenin were manifest in patient BM-MNC (n=6)compared to normal donors (n=3), as well as in normal BM-MNC treatedwith 5 μg/mL rhS100A9 for 24 hours compared to controls (p=0.043 andp=6.38×10⁻⁷, respectively) (FIG. 6c, 6d ). Finally, β-catenin geneexpression was increased 9.5-fold in the BM cells of S100A9Tg miceversus WT BM cells, with corresponding up-regulation of Wnt/β-catenintarget genes. Thus, S100A9-directed activation of β-catenin is ahallmark of MDS.

To test if somatic gene mutations manifest in MDS trigger pyroptosis andenhance self-renewal via activation of β-catenin, this circuit was firstinvestigated in cells engineered to express MDS-associated mutants ofthe U2AF1 splicing factors (Yoshida K, et al. Nature. 2011478(7367):64-9). The percentage of pyroptotic cells was increased4.6-fold in S34F U2AF1 mutant-versus WT U2AF1-expressing cells,accompanied by increased levels of a-caspase-1 (p=0.044) and annexin-V(p=0.021) (FIG. 15a-15h ). U2AF1-S34F-expressing cells also displayedsignificant increases in mean cell area (p=0.035) and ethidium bromideinflux (FIG. 15i, 15j ), the mean percentage of ROS⁺ cells (p=1.5×10⁻³),ROS MFI (p=0.032) (FIG. 6e, 6f ) and nuclear localization of β-catenin(FIG. 6g, 6h ). Notably, treatment of U2AF1-S34F mutant cells with theanti-oxidant N-acetylcysteine (NAC) or the NADPH oxidase (NOX) inhibitorDPI effectively reduced β-catenin activation in mutant cells (p=3.8×10⁻³and p=2.5×10⁻⁶, respectively) (FIG. 6g, 6h ).

BM cells harvested from SF3B1-K700E conditional knock-in mice (n=3),which also display a MDS phenotype (Obeng E. A, et al. Blood. 2014124(6):828-30), displayed similar increases in the percentage ofpyroptotic versus apoptotic cells, with significant increases in totala-caspase-1⁺ cells (p=0.014) versus WT controls (n=3) (FIG. 16a, 16b ).Further, MFI of a-caspase-1 and a-caspase-3 were both significantlyincreased in the SF3B1-K700E mutant BM cells (p=0.030 and p=6.92×10⁻³,respectively) (FIG. 16c ), which displayed inflammasome assembly (FIG.16d ). Accordingly, NLRP3 protein expression was increased 1.9-fold inthe SF3B1-K700E cells (p=0.063), as NLRP3 inflammasome formation wasincreased 1.2-fold (FIG. 16e ). Inflammasome formation was also assessedby flow cytometry based on the detection of the inflammasome adaptorprotein ASC oligomerization, whose incorporation into inflammasomecomplexes can be detected by changes in fluorescence pulse height andarea (Sester D. P, et al. J Immunol. 2015 194(1):455-62). SF3B1-K700Emutant-expressing BM cells (n=6) had marked increases in inflammasomeformation versus WT controls (n=6, p=8.4×10⁻³), which was significantlyreduced upon treatment with NAC (n=6, p=2.68×10⁻³) or DPI (n=3) (FIG.16f, 16g ). Moreover, mean percentage of ROS⁺ cells and ROS MFI wereboth markedly increased in SF3B1-K700E-expressing mutant BM cells, whichwas extinguished below the level of WT controls by NAC or DPI treatment(FIG. 17a, 17b ). Finally, SF3B1-K700E mutant-expressing BM cellsdemonstrated a statistically significant increase in the percentage ofcells with elevated levels of nuclear β-catenin compared to WT controls(p=0.40), which was significantly reduced upon treatment with NAC(p=2.0×10⁻³) or DPI (p=1.8×10⁻²) (FIG. 17c, 17d ). Similar findings wereobserved in mutant versus WT SRSF2-expressing HEK293T cells (FIG. 18),as well as with epigenetic regulatory gene mutations (ASXL1, TET2) (FIG.19, 20). Thus, MDS somatic gene mutations prime cells to undergopyroptosis, which promotes self-renewal and contributes to aninflammatory microenvironment that is driven by ROS.

DISCUSSION

Heretofore ineffective hematopoiesis in MDS has been attributed to highfractions of proliferating BM progenitors with a propensity to undergoapoptotic cell death within an unexplained inflammatory microenvironment(Span L. F, et al. Leuk Res. 2007 31(12):1659-67; Raza A, et al. Blood.1995 86(1):268-76). Nearly two decades ago it was reported that MDS HSPCgenerate IL-1β in short term cultures, which directly correlated withthe extent of apoptosis as measured by DNA fragmentation (Mundle S. D,et al. Blood. 1996 88(7):2640-7). Evidence is disclosed that these andother biological features of MDS are explained by the activation of theNLRP3 pattern recognition receptor by S100A9 and by ROS DAMPintermediates that induce inflammasome assembly, β-catenin nucleartranslocation and pyroptotic cell death. Notably, pyroptotic-associatedgene transcripts and inflammasome assembly are profoundly up-regulatedin MDS independent of genotype. Moreover, pyroptotic but not apoptoticcells are markedly increased in MDS stem cells, progenitors, anderythroid precursors. Accordingly, knockdown of caspase-1, but notcaspase-3, in MDS BM-MNC, significantly reduced the pyroptotic cellfraction. Similarly, neutralization of S100A9 in MDS BM plasma, orpharmacologic inhibition of inflammasome assembly, suppressed pyroptosisand restored effective hematopoiesis in vitro and in a murine MDS model.Thus, pyroptosis, a caspase-1-dependent inflammatory cell death, impairsHSPC survival in MDS.

S100A8/S100A9 activate both NF-κB and NLRP3 inflammasome assembly via anNADPH oxidase (NOX)/ROS-dependent mechanism (Simard J. C, et al. PLoSOne. 2013 8(8):e721381; Liao P. C, et al. Inflamm Res. 2013 62(1):89-96;Heid M. E, et al. J Immunol. 2013 191(10):5230-8; Bauernfeind F, et al.J Immunol. 2011 187(2):613-7). S100A8/9 heterodimers serve as a scaffoldfor the membrane assembly and activation of the NOX complex (DoussiereJ, et al. Eur J Biochem. 2002 269(13):3246-55; Kerkhoff C, et al. FASEBJ. 2005 19(3):467-9), which generates ROS via transfer of electronsacross membranes to generate superoxide (Bedard K, et al. Physiol Rev.2007 87(1):245-313). NOX activity regulates both priming and activationof NLRP3 inflammasomes, as NOX inhibition suppresses the activation ofcaspase-1 and IL-1β secretion (Liao P. C, et al. Inflamm Res. 201362(1):89-96). Moreover, transcription and nuclear localization ofβ-catenin are redox- and NOX1-dependent (Coant N, et al. Mol Cell Biol.2010 30(11):2636-50; Wu X, et al. Cell. 2008 133(2):340-53). AlthoughMDSC are a key paracrine source of S100A9 in the MDS BM-microenvironment(Chen X, et al. J Clin Invest. 2013 123(11):4595-611), here it shownthat MDS HSPC also express high intracellular levels of S100A9 (FIG.11), suggesting that inflammasome activation may be sustained byintracrine DAMP stimulation, and upon cell lysis, promote BM expansionof MDSC. Importantly, NOX inhibition suppressed inflammasome andpi-catenin activation in both patient-derived BM-MNC and cells harboringvaried classes and types of MDS founder gene mutations. Thus, S100A9induces NOX activity to drive ROS-dependent inflammasome assembly andpyroptosis, accompanied by β-catenin nuclear translocation.

A hallmark of MDS BM precursors is their enlarged cell size ormacrocytosis. It is shown that activation of pattern recognitionreceptors triggers expansion in size of MDS progenitors via influx ofcations mediated by the transient receptor potential melastatin 2(TRMP2) cation channel, a plasma membrane calcium-permeable channel inhematopoietic cells (Zhang W, et al. Am J Physiol Cell Physiol. 2006290(4):C1146-59). Further, TRPM2 channels are activated by NOX-derivedROS via oxidation of a single channel methionine residue, Met-214, whichis indispensable for NLRP3 inflammasome activation (Kashio M, et al.Proc Natl Acad Sci USA. 2012 109(17):6745-50; Zhong Z, et al. NatCommun. 2013 4:1611; Yamamoto S, et al. Nat Med. 2008 14(7):738-47).Activation of TRPM2 then directs calcium influx that then leads tocorresponding increases in cell volume (Kuhn F, et al. Pflugers Arch.2005 451(1):212-9). The disclosed data show that MDS BM-MNC displayincreased influx of the TRPM2 channel substrate ethidium bromide,confirming inflammasome-initiated pore formation. Additionally,quantifying BM cell size according to lineage and stage of maturationconfirmed the larger size of MDS BM precursors versus normal controls.The disclosed findings indicate that S100A9-mediated NOX activation andsubsequent inflammasome initiated pyroptosis explain the characteristiclarger cell size, proliferation and inflammatory cell death manifest inMDS.

NOX-derived ROS enhance mitogenic response to receptor-tyrosine kinasesthrough oxidative inactivation of protein tyrosine phosphatases (Block K& Gorin Y. Nat Rev Cancer. 2012 12(9):627-37). Somatic gene mutationsfound in MDS are known to trigger Rac1/NOX-dependent ROS generation(Sallmyr A, et al. Cancer Lett. 2008 270(1):1-9; Rassool F, et al.Cancer Res. 2007 67(18):8762-71). As ROS serve as DAMP intermediatesthat activate both inflammasomes and β-catenin, ROS generated by eitherS100A9 or MDS somatic gene mutations may drive pyroptosis, self-renewaland propagation of the MDS clone. Mechanistically, NOX-derived ROSstabilizes and activates β-catenin by oxidation and dissociation ofnucleoredoxin (NRX) from disheveled (Dvl), which in turn inactivates theβ-catenin destruction complex (Funato Y, et al. Nat Cell Biol. 20068(5):501-8). It is shown herein that ROS and nuclear β-cateninlocalization are profoundly increased in MDS HSPC. Further, S100A9treatment of normal BM-MNC is sufficient to trigger nucleartranslocation of β-catenin that is abolished by the anti-oxidant NAC orNADPH-oxidase inhibition. Similarly, BM-MNC from S100A9-Tg micedisplayed marked increases in the expression and nuclear localization ofβ-catenin, with corresponding up-regulation of β-catenin target genes(data not shown). Of particular importance, varied RNA splicing genemutations (U2AF1, SF3B1, SRSF2) and epigenetic regulatory gene mutations(ASXL1, TET2) found in MDS triggered pyroptosis, pore formation, cellvolume expansion and β-catenin activation, which was extinguished bytreatment with NAC or NOX-inhibition. Thus, both S100A9-induced NOXactivation and MDS gene mutations initiate pyroptosis through superoxidegeneration to drive β-catenin activation and afford a proliferativeadvantage to the MDS clone. Accordingly, these findings explain how suchdiverse somatic gene mutations give rise to an MDS phenotype.

In conclusion, despite genetic heterogeneity, inflammasome activationunderlies the biological phenotype in MDS, whereby DAMP signals and MDSgene mutations license a common redox-sensitive inflammasome platform todrive pyroptotic death, elaborate inflammatory cytokines, activatecation influx, and support propagation of the MDS clone throughβ-catenin activation (FIG. 21). These findings provide a common platformthat accounts for the biological features of MDS and suggest thatstrategies targeting S100A9 neutralization or inhibition of pyroptosissignaling offer therapeutic promise in MDS.

Example 2: Pyroptotic ASC Specks as a Diagnostic Biomarker inMyelodysplastic Syndromes

Methods

PB plasma samples were collected from normal donors (n=18) and patientswith MDS (n=45), AML (n=10), CMML (n=20) and CLL (n=50), which providedgreater than 90% power to detect significant differences in bothpercentage and mean fluorescent intensity (MFI) of ASC specks. 300micrograms of protein was aliquoted from each donor and stained withanti-ASC antibodies. Sample acquisitions were made on a BD FACSCaliburflow cytometer.

Results

Percentage (FIG. 22A) and MFI (FIG. 22B) ofASC specks were significantlygreater in the PB (n=45) than in the BM (n=47) of MDS patients(p=2.0×10⁻³ and 3.5×10⁻³, respectively), prompting further investigationof PB specks. No significant difference in speck percentage or MFI wasdetected in patients of lower versus higher-risk MDS, suggestingdiagnostic and not prognostic use. Mean percentage of ASC specks andspeck MFI was significantly greater in MDS compared to normal donors(p=1.7×10−3 and 0.010, respectively), whereas normal donor levels didnot differ from other hematologic malignancies studied. MDS samplesdisplayed significantly greater percentage and MFI of specks compared topatients with AML (p=1.4×10⁻¹³ and 3.2×10⁻⁹, respectively), CMML(p=1.2×10⁻¹⁰ and 1.0×10⁻⁷, respectively) and CLL (p=9.9×10⁻¹¹ and3.3×10⁻⁷, respectively).

Unless defined otherwise, all technical and scientific terms used hereinhave the same meanings as commonly understood by one of skill in the artto which the disclosed invention belongs. Publications cited herein andthe materials for which they are cited are specifically incorporated byreference.

Those skilled in the art will recognize, or be able to ascertain usingno more than routine experimentation, many equivalents to the specificembodiments of the invention described herein. Such equivalents areintended to be encompassed by the following claims.

What is claimed is:
 1. A method for treating a myelodysplastic syndrome(MDS) in a subject suspected of having a hematological disorder,comprising assaying a sample from the subject to detect inflammasomeactivation, wherein an increase in inflammasome activation in the samplecompared to a control is an indication of MDS in the subject, andtreating the subject with a therapeutically effective amount of aninflammasome inhibitor.
 2. The method of claim 1, wherein the samplecomprises hematopoietic stem/progenitor cells (HSPC).
 3. The method ofclaim 1, wherein the sample comprises a bone marrow sample.
 4. Themethod of claim 1, wherein the inflammasome comprises an NLRP3inflammasome.
 5. The method of claim 1, wherein inflammasome activationis detected by detecting apoptosis associated speck-like proteincontaining a CARD (ASC) specks.
 6. The method of claim 1, whereininflammasome activation is detected by detecting CASP-1 gene expression.7. The method of claim 1, wherein inflammasome activation is detected bydetecting NLRP3 gene expression.
 8. The method of claim 1, whereininflammasome activation is detected by detecting active caspase-1protein levels.
 9. The method of claim 1, wherein inflammasomeactivation is detected by detecting NLRP3 protein levels.
 10. The methodof claim 9, wherein inflammasome activation is detected by detectingco-localization of active caspase-1 and NLRP3 protein.
 11. The method ofclaim 1, wherein inflammasome activation is detected by detecting NLRP3inflammasome complexes.
 12. The method of claim 1, wherein inflammasomeactivation is detected by detecting IL-1β levels, IL-18, levels, or acombination thereof.
 13. The method of claim 1, further comprisingpredicting from the detection of inflammasome activation whether thesubject has low-risk or high-risk MDS.